Bacterial, Mycoplasmal, and Mycotic Infections

C H A P T E R

12 Bacterial, Mycoplasmal, and Mycotic Infections Steven H. Weisbroth1, Dennis F. Kohn2 1

Consultant, Laboratory Animal Medicine, St. John’s University, McLean, VA, United States; 2Professor Emeritus, Pathology, Columbia University, New York, NH, United States

I. INTRODUCTION This chapter reviews the microbiologic, pathologic, diagnostic, and medical features of the pertinent bacterial, mycoplasmal, and mycotic (BMM) infectious diseases of the laboratory rat. The issue of pertinence is important because of the changing prevalence of the agents under discussion. Nonetheless, the agents considered in this review will adhere closely to the harmonized bioexclusion profiles recommended by the Federation of European Laboratory Animal Science Association (FELASA) and American Association forLaboratory Animal Science (AALAS) (Table 12.1) and which are generally recognized globally by animal health experts, rodent producers, and scientific users as necessary components of rat health surveillance programs (Guillen, 2012). From the historical perspective it is important to recognize that an evolutionary process has been at work on the microbiomes of the laboratory rat since its domestication more than a century ago (Lindsey and Baker, 2005; Robinson, 1965). Because the ecological niche of the laboratory rat has been sequentially modified by refinements in both rodent breeder production methodology and user laboratory environments, the pathogenic microbial species accompanying these changes have shifted to forms largely unrecognized as rat pathogens at the beginning of the process. As a general statement it may be said that the primary microbial pathogens causing major epizootic (and zoonotic) hazards were among the first to be diminished or eradicated by these ecological changes. Examples include Yersinia sp., Salmonella sp., Leptospira sp., and Streptobacillus moniliformis, which figured prominently as agents encountered in the decades just before and after the turn of the 20th century. Under the generally improved sanitary standards of laboratory animal care environments obtaining between the 1900s and the 1950s, the main

The Laboratory Rat, Third Edition https://doi.org/10.1016/B978-0-12-814338-4.00012-X

accomplishments of disease control programs were to reduce the incidence of primary pathogens and disease outbreaks. By the 1950s, as a practical matter, clinically apparent disease caused by BMM had been reduced to a small number of extremely important agents of high morbidity that nonetheless imposed substantial limitations on the ability to conduct research with rats in the absence of variables imposed by disease. The microbial components of this spectrum were now recognized to include Mycoplasma pulmonis, Streptococcus pneumoniae, Clostridium piliforme, Corynebacterium kutscheri, and Pasteurella pneumotropica, entities of minor or unrecognized significance at the turn of the 20th century. Recognition of the deleterious effects of disease on the research process did not at that time include viral agents of rats. The key scientific breakthroughs to reduce or eliminate these pathogens were techniques to exclude agents of disease by gnotobiotic derivation (Foster, 1959; Carter and Foster, 2005). This process, described fully in Chapter 21, when employed in large-scale commercial rodent production, enabled common availability of rats essentially free of pathogenic BMM to the scientific community in the 1970s, and later, BMM and virus-free rats in the 1980s and 1990s (Foster and Pfau, 1963). While it is true that all pathogens may be excluded from rats held within gnotobiotic isolators, once removed to production barriers or to user facilities, a lesser degree of shielding obtains and the microbiome diversity of the host becomes variably enlarged by environmental and human contacts and other institutional rodent residents, thus opening up potential for infection by opportunistic commensals, and to a lesser degree primary pathogens. This point has been noted previously (Boot et al., 1989; ILAR, 1998). In fact, the potential infection of incoming pathogen-free rodent stocks by resident institutional populations carrying acquired microbial pathogens has been a cause of ongoing concern (Barthold, 2008).

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TABLE 12.1 FELASA and AALAS Recommended Bioexclusion Profiles of Bacterial, Mycoplasmal, and Fungal Pathogens for Monitoring the Health Status of Colonies and Transported Cohorts of Laboratory Rats: Changes Over Time. FELASA (1994)a

FELASA (1996)b

FELASA 2002c

FELASA (2014)d

FELASA/AALAS (2014)e

Mycoplasma sp. Pasteurella sp. Salmonella spp. Streptobacillus moniliformis Streptococcus b-hemolytic Streptococcus pneumoniae Corynebacterium kutscheri Bordetella bronchiseptica Leptospira spp. If indicated/Optional CAR bacillus Dermatophytes Escherichia coli Klebsiella pneumoniae Klebsiella oxytoca Pneumocystis spp. Proteus spp. Pseudomonas aeruginosa Staphylococcus aureus Yersinia pseudotuberculosis

Mycoplasma sp. Pasteurella sp. Salmonella spp. Streptobacillus moniliformis Streptococcus b-hemolytic Streptococcus pneumoniae Corynebacterium kutscheri Bordetella bronchiseptica Clostridium piliforme Leptospira spp. Citrobacter freundii 4280 If indicated/Optional not enumerated

Mycoplasma sp. Pasteurellaceae Salmonella spp. Streptobacillus moniliformis Streptococcus b-hemolytic Streptococcus pneumoniae Corynebacterium kutscheri Bordetella bronchiseptica Clostridium piliforme Helicobacter spp. Pneumocystis spp. If indicated/Optional not enumerated

Mycoplasma pulmonis Pasteurella pneumotropica Salmonella spp. Streptobacillus moniliformis Streptococcus b-hemolytic Streptococcus pneumoniae Clostridium piliforme Helicobacter spp. Pneumocystis spp. If indicated/Optional Bordetella bronchioseptica Corynebacterium kutscheri Klebsiella pneumoniae Klebsiella oxytoca Other Pasteurellaceae Pseudomonas aeruginosa Staphylococcus aureus Encephalitozoon cuniculi

Mycoplasma pulmonis Pasteurella pneumotropica Salmonella spp. Streptobacillus moniliformis Streptococcus b-hemolytic Streptococcus pneumoniae Corynebacterium kutscheri Bordetella bronchioseptica Clostridium piliforme Helicobacter spp. Pneumocystis spp. CAR bacillus Bordetella hinzi Klebsiella pneumoniae Klebsiella oxytoca Pseudomonas aeruginosa Staphylococcus aureus

a

Kraft et al. (1994) Laboratory Animals 28: 1e12. Rehbinder et al. (1996) Laboratory Animals 30: 193e208. c Nicklas et al. (2002) Laboratory Animals 36: 20e42. d Mahler et al. (2014) Laboratory Animals 48: 178e192. e Pritchett-Corning et al. (2014) J Am Assoc Lab Anim Sci 53: 633e640. f FELASA=Federation for Laboratory Animal Science Association. g AALAS=American Association for Laboratory Animal Science. b

With reference to Table 12.1, it will be seen that over the years since the initiation of formal rodent health surveillance programs there is a core of some 8e10 indigenous BMM agents that are regarded, depending on circumstances, as primary pathogens associated with infectious disease. These agents include M. pulmonis, P. pneumotropica, Salmonella sp., S. moniliformis, S. pneumoniae, b-hemolytic streptococcus sp., C. kutscheri, C. piliforme, Helicobacter sp., and Pneumocystis carinii. Most, but not all, of the agents listed here as primary pathogens share the pathogenic capability of being able to fulfill Koch’s third postulate, which states that to be designated as causative, isolates from a diseased animal must be capable of causing a similar disease when used to challenge healthy animals (Hornef, 2015; Hoffman et al., 2016). With minor exceptions, the issue of pertinence bears on the point that in the 10e12 years since the second edition of this text, none of the agents included here as primary pathogens have been reported in the literature as a cause of clinical disease in laboratory rats; a telling indicator of diminished significance. The issue of contemporaneous prevalence in laboratory rats is important as well, but difficult to put dimensions on because of infrequent, irregular, and fragmentary reporting. Such evidence as exists derives from reports of institutional health surveillance screens that indicate that some of the primary pathogens may still be

encountered as inapparent infections (Mahler and Kohl, 2009; Zeriner and Regault, 2003; Carty, 2008; Jacoby and Lindsey 1997, 1998; Schoondermark-van de Ven et al., 2003; Pritchett-Corning et al., 2009; Liang et al., 2009). Moreover, it should not be overlooked that these agents persist in wild, fancy, and pet rat populations and in conventional research rat residents, and also that there are other regions in the research world where the pathogen status of research rodents cannot be assumed (Otto et al., 2015), thus the potential for ingress to pathogen-free stocks remains. A range of other BMM, many of which are also listed in Table 12.1, are variably described as “weakly pathogenic” or “opportunistic,” and are thought of as having lesser ability to cause disease than the “primary pathogens.” These agents are usually clinically inapparent and typically unable to meet Koch’s third postulate. Such microbial forms as C. kutscheri, P. pneumotropica, P. carinii, C. piliforme, Clostridium difficile, S. moniliformis, and Pseudomonas aeruginosa characteristically occur in immunocompetent rats as inapparent infections. They are regarded as opportunists and occasionally encountered as causes of disease in laboratory rats, usually in situations where innate host resistance has been compromised by genomic immunodeficiency or other heritable constitutional factors or by exogenous physiologic stressors that transiently impair resistance such as

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shipment and relocation or when coupled with other intercurrent infections. It is also recognized that research treatments that vitiate innate resistance or immunocompetence, e.g., vitamin deficiency, nuclear irradiation, administration of adrenal corticosteroids and other immunosuppressive drugs, and thymectomy, are known examples of the types of experimentally induced precedent conditions known to trigger microbial escape from inhibition by invasive opportunistic pathogens. It is necessary at this point to briefly introduce the concept of the rodent microbiome and how it is increasingly being recognized as regulating both the colonization and invasiveness of microbial agents having pathogenic potential. It is an area of intense interest at present because of the newly unfolding understanding of the profound effects of the microbiome on mammalian health, nutrition, and physiology. The microbiome may be defined as the aggregate of the 500e1000 microbial species inclusive of commensals, symbionts, opportunists, pathobionts, and pathogens that make up the microbial population of the mammalian body (Bleich and Fox, 2015). The diversity of species composing the microbiomes of the various body locations (oral cavity, gastrointestinal, nasopharyngeal, skin, and urogenital) varies depending on the cultural milieu provided by the several locations and required by the respective microbes. There are several recognized means by which the autochthonous commensal, symbiont, and pathobiont components of the microbiome act to inhibit populations or limit invasiveness of opportunists (and pathogens). These mechanisms include microbial competition for nutrients and niche space, effecting pH conditions inhibitory to opportunists, enhancement of host defense mechanisms mediated by commensal microbes, e.g., by stimulation of mucosal irritation and heightened reactive immunological responsiveness, and production of exogenous factors directly inhibitory to certain pathogens or which prevent the development of pathogenic properties, e.g., capsules and still others incompletely known but under active investigation (Kamada et al., 2013; Bleich and Fox, 2015; Hoffman et al., 2016; Piters et al., 2016; and Chan et al., 2016). It should be understood that the gradient of microbial species making up the microbiomes of wild and feral rats is assumed to be more diverse than that of domesticated pet or fancy rats (unfortunately there is no comparative information on the subject), more so than that of conventionally bred or conventionally maintained rats at user institutions, much more diverse than that of rats in gnotobiotically derived breeding barriers and of accreditably maintained rats at user institutions, and considerably less restricted, of course, than that harbored by Schaedler flora-associated and germ-free gnotobiotes housed in isolators (Schaedler and Orcutt, 1983; Orcutt et al., 1987). Furthermore, as the microbiome of the laboratory rat

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continues to narrow over time under the more limited microbial diversity permitted by the summative total of gnotobiotic rederivation procedures and microbiologic housing barriers imposed by breeders and users alike, the inhibitory functionality of the microbiome is correspondingly lessened. In fact, the progressively more restricted commensal, symbiont, and pathobiont forms making up the microbiomes of gnotobiotically derived rodents would seem to paradoxically underlie the increasing incidence and diversity of case reports of newly recognized opportunistic microbial species. As the restricted microbial diversity imposed by gnotobiotic derivation proceeds from the breeding barrier to user facilities, the potential sources for adding to microbiome diversity, including potential pathogens, are essentially limited to those of (1) chance exposures from other, variably restricted, institutional rodent residents, (2) the regulated animal care environment, and (3) the animals’ human contacts. These are the sources for the opportunists encountered in recent years (ergo Pogo: we have met the enemy and he is us!). These factors prevent universally applicable guidelines to usefully categorize colonizing microbes as “primary pathogens,”“opportunists,” or even as “commensal” flora because it is recognized that the ability to cause disease is determined as much by constitutional factors of susceptibility and resistance of the host and its interaction with the microbial regulating properties of the regional microbiomes, as it is by microbial factors of pathogenicity. It can be anticipated that as time goes on, many of the primary pathogens in the core list will be recognized as having been essentially eradicated from the structured world of the laboratory rat. They will be relegated to the status as being of only historical significance and either dropped from health surveillance microbial profiles or tested for with diminished frequency (as several have been already, see Table 12.1). Recognition of this point has already shaped health surveillance programs at many institutions resulting in risk-based allocation of surveillance resources based on (presumptive) pathogen prevalence (Pritchett-Corning et al., 2009). At the same time, the diversity of microbial forms conventionally regarded as “opportunistic” or even as “commensal” that may now be seen as causes of disease in certain circumstances will surely increase. Thus the list of microbial pathogens of the laboratory rat should not be regarded as a fixed list, but rather as having a forward-moving boundary in which old pathogens are eradicated and newly recognized causes of disease emerge, effecting periodic reconstitution of the bioexclusion profiles (Weisbroth, 1999a; Weisbroth et al., 2006). We can see this point taking place over time in Table 12.1. This final section brings to readers’ attention a number of recent developments that improve detection and

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identification of BMM on the FELASA exclusion list, as well as viral and parasitic pathogens. These technical developments include: 1. The advent of molecular methodology for structuring microbial genetic relationships has led to numerous changes with reclassification and renaming of many microorganisms and delineation of many new species. Readers’ attention is drawn to Table 12.2, which summarizes nomenclature changes as they pertain to the agents in this chapter. Faster and more accurate identification of isolated microbes by the technique of matrix-assisted laser desorption/ ionization time-of-flight mass spectrophotometry (MALDI-TOF MS) has enabled rapid recognition and discrimination between closely related forms (Seng et al., 2009). The principle is to match determined peptide spectral patterns of the isolated microbe against an open database to identify the clinical isolate down to the species level. Use of MALDI-TOF MS represents a growing alternative to traditional phenotypic markers, i.e., biochemical, cultural, and morphologic criteria and polymerase chain reaction (PCR) identification protocols (Dieckmann and Malony, 2011; Eisenberg et al., 2015a,b; 2016a,b; Goto et al., 2012; Harju et al., 2017; Kuhnert et al., 2012; Michel et al., 2018; Packeu et al., 2013; Werno et al., 2012). In addition to accuracy, the advantages of MALDI-TOF MS include speed, on the order of minutes or hours rather than days or weeks, and the low cost of individual assays. It is also true that the apparatus is expensive and its use requires specific expertise. 2. Advances in solid-phase serologic technology, where antigens are adsorbed onto some form of solid structure where the antigeneantibody reaction occurs, have enabled miniaturization of reaction platforms and use of multiplexed antigenic arrays within a single well, tube, chip, or suspension of polystyrene beads (Shek et al., 2015). Miniaturization of the platform allows utilization of as little as 1e2 m of sera as sufficient to test against an array of microbial antigens (as may be eluted from a single drop of dried blood, see 4. ahead). In these indirect tests, the titer of antibody is directly related to the intensity of a measurable signal, e.g., fluorescence. The separate antigeneantibody reactions can occur simultaneously and be spectrophotometrically analyzed for intensity, thus facilitating high throughput. In principle, any agent for which a directly derived or recombinant antigen can be developed can be used to coat solid-phase carrier platforms. Such antigen arrays are called multiplex (Khan et al., 2005; Richens et al., 2010; Wunderlich et al., 2011) and acronyms have come into common

usage to denote these platforms, e.g., median fluorescence intensity (MPI) and multiplexed fluorometric immunoassay (MFIA). Use of multiplexed formats have raised the efficiency and economical testing of rodent sera from sentinel rodents, for independently validating rodent producer health data, for investigating diagnostic sera from disease outbreaks, and for detection of antibodies from test rodents inoculated with rodentorigin biologics and tissue cultures potentially contaminated with rodent pathogens (Mouse Antibody Production [MAP] and Rat Antibody Production [RAP] tests) (Adams and Myles, 2013). 3. Recombinant antigens, mentioned earlier, have been developed for a number of rodent pathogens that are zoonotically dangerous in viable mode, or which are fastidious, difficult, or uncultivable to propagate for the purpose of preparing test antigens with the purity, concentration, and volume needed for solid-phase serologic platforms. In principle, a cloned gene inserted into a microbial or baculovirus (insect virus) vector can be used to produce large quantities of recombinant protein antigen that is noninfectious and can be purified to meet various purposes. These antigens can be used to coat a solid platform, e.g., wells or test beads to assemble pathogen detection panels of enhanced inclusivity and accuracy (Shek et al., 2015). 4. Dried blood spot (DBS) technology has been facilitated by MFIA multiplexed formats that use very small volumes of sera, as may be obtained by elution of the sera from a drop of blood dried onto a paper disc or card. This alternative to preparation of a serum sample has a number of advantages: collection is minimally invasive and may be obtained from unanesthetized animals, e.g., sentinels without their removal from test sites. So doing enables serial testing of the same sentinels over time, thus conserving animal usage. Use of the DBS alternative eliminates the steps needed to prepare diluted serum samples for submission and DBS can be transported or shipped in an envelope at ambient temperature without coolant or bulky insulation (Smit et al., 2014). 5. PCR technology for diagnostic purposes has improved as well in recent years. Use of the TaqMan assay employs a fluorescent nuclease where the intensity of the fluorescent signal is determined spectrophotometrically at each amplification cycle as it occurs, i.e., in real time thus allowing quantification of the signal and a means of discriminating between higher concentrations of the signal(s) of interest, e.g., the genomic sequence of a pathogen in the sample, from lower-level background or insignificant amplification (Shek et al., 2015). Real-time PCR assays are denoted as quantitative PCR (qPCR). Use of the TaqMan assay in an open array format enables multiplex separate

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I. INTRODUCTION

TABLE 12.2

Changes in Microbial Nomenclature.

Older Nomenclature

Newer or Reclassified Nomenclature

Streptococcus, fecal, l-hemolytic

Enterococcus ratti

Bacillus piliformis

Clostridium piliforme

Rules for nomenclature designations of Salmonella species: (Refs. 1e2) 1. Serotypes (serovars) are not italicized. The first letter is capitalized. 2. Serotypes (serovars) do not have standing in nomenclature. 3. After first use, nomenclature can be shortened, namely Salmonella enterica serovar Typhimurium can be shortened to Salmonella Typhimurium. Rodent Salmonella species

Salmonella enterica subspecies enterica

Salmonella typhimurium

Salmonella enterica serovar Typhimurium

Salmonella typhi

Salmonella enterica serovar Typhi

Salmonella cholerasuis

Salmonella enterica serovar Cholerasuis

Salmonella enteriditis

Salmonella enterica serovar Enteriditis

Salmonella galinarum

Salmonella enterica serovar Galinarum

Salmonella paratyphi A

Salmonella enterica serovar Paratyphi A

Salmonella paratyphi B

Salmonella enterica serovar Paratyphi B

Salmonella salamae

Salmonella enterica serovar Salamae

Other Salmonella species of potential interest

Salmonellaenterica subspecies arizonae Salmonellaenterica subspecies diarizonae Salmonellaenterica subspecies houtenae Salmonellaenterica subspecies indica

Pasteurellaceae (in which rat is the primary host, except for Muribacter muris) (Refs. 3e5) Pasteurella pneumotropica, biotype Jawetz

Rodentibacter pneumotropicus

Pasteurella pneumotropica, biotype Heyl

Rodentibacter heylii

Pasteurella pneumotropica,

Rodentibacter ratti

Taxon B [V-factor dependent] Taxon 22 Taxon 17

Rodentibacter rarus

Haemophilus(V-factor dependent)

Rodentibacter heidelbergensis

Haemophilus(V-factor dependent)

Rodentibacter trehalosifermentans

Actinobacillus muris

Muribacter muris

CAR bacillus

Filobacterium rodentium (Ref. 6)

Haemobartonella muris

Mycoplasma haemomuris (Ref. 7)

Encephalitozoan cuniculi (had been classified in Kingdom Protozoa, now classified in KingdomFungi) (Ref. 8) Pneumocystis

(Refs 9e10)

Rat Pneumocystis, most common

Pneumocystis carinii Pneumocystis wakefieldiae

Rat Pneumocystis, least common

Pneumocystis carinii f. sp. rattus-secundi Pneumocystis carinii f. sp. rattus-tertii Pneumocystis carinii f. sp. rattus-quarti Continued

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Changes in Microbial Nomenclature.dcont’d

Mouse Pneumocystis

Pneumocystis murina

Rabbit Pneumocystis

Pneumocystis oryctolagi

Human Pneumocystis

Pneumocystis jirovecii

1

Tindall (2005). Fookes et al. (2011). Adhikary et al.(2017a,b). 4 Boot et al. (2018). 5 Nicklas et al. (2015). 6 Ike et al. (2016). 7 Neimark et al. (2002). 8 Capella-Gutierrez et al. (2012). 9 Palmer et al. (1999). 10 Palmer et al. (2000). 2 3

analysis for the presence of each sequence of interest in the sample. These formats have been deployed to allow simultaneous qPCR testing for the presence of virtually all agents in the FELASA exclusion panels in Table 12.1 (Henderson et al., 2013). 6. Recent improvements in the testing of the animal care environment have also been developed. Real-time (qPCR) assays are used to detect the presence of a range of rodent pathogens collected by swab from any of the surfaces in the animal care environment, including ventilated caging, quarantine, and holding room exhaust air duct (EAD) manifolds and plenums. In practice, for economy and efficiency, swabs can be pooled, and dust contents eluted and tested for any of a number of sequences of interest. In controlled field studies, EAD test swabs were used to detect a number of pathogens from rodents in ventilated cages (P. pneumotropica, Helicobacter spp., and several viruses and parasites), but did not detect norovirus or mouse parvovirus (Bauer et al., 2016). Several of these agents (P. pneumotropica, Helicobacter spp., and fur mites) were often known to be undetectable in samples taken from soiled bedding sentinels, but detected as present in the same population monitored by EAD samples (Jensen et al., 2013; Manuel et al., 2016; Miller et al., 2016). In summation on these advances in diagnostic technology, they have had a transformative effect on the process of rodent health surveillance, including all of the agents in the FELASA exclusion panels. For reasons of economy, efficiency, and accuracy, the taking of traditional diagnostic necropsy samples from representative lots of animals, whether derived from institutional sentinels or from breeder production units, can be obviated by some combination of the advances outlined earlier. For example, the combination of a serum or DBS sample for serology, rectal, fur, and oropharyngeal swabs for PCR and EAD swabs for PCR can be used to effectively determine the presence or valid absence of all of the agents in the FELASA exclusion lists. It is also true, however, that no single test modality can yield all of the needed information about pathogen

status of the larger group represented by the test samples. Moreover, each of the modalities is accompanied by certain limitations, caveats, and qualifications of timing and process that are detailed in the chapter by Shek et al. (2015). Thus professional judgment, institutional needs, and budget should determine the healthmonitoring test parameters.

II. BACTERIAL INFECTIONS A. Streptobacillus moniliformis (Streptobacillosis) Considered only within the context of its status as a pathogen of laboratory rats, S. moniliformis occupies a somewhat ambiguous role. It has been circumstantially associated with various aspects of respiratory disease in laboratory rats since early times; however, two other features of the organism have received intense scrutiny, tending to obscure or deemphasize the limited potential for pathogenicity of the organism in its natural reservoir host. These other features include the etiological significance of S. moniliformis in rat bite and Haverhill fevers of humans, and the cultural instability of the organism leading to L-phase variants that for a certain period of time were confused with Mycoplasma of rat origin. Taxonomically, the agent is positioned in Class Fusobacteria, Order Fusobacteriales, Family Leptotrichiaceae, with Genus and species as S. moniliformis. Despite its monospecific status for almost 90 years, the genus is now recognized has holding an additional number of species, including Streptobacillus hongkongensis, and Streptobacillus felis based on genomic analysis (Eisenberg et al., 2015a,b), several recently characterized species, including Streptobacillus ratti characterized from black or house rats (Rattus rattus) (Eisenberg et al., 2016a,b), and Streptobacillus notomytis isolated from the hopping mouse, Notomys alexis (Eisenberg et al., 2015a,b). The Norway rat is recognized as the preeminent reservoir host for S. moniliformis and is carried asymptomatically by most

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wild Rattus norvegicus, and quite commonly by pet and fancy rats. S. moniliformis has been essentially eradicated from laboratory rats by the gnotobiotic process, but is still encountered serologically or by PCR during diagnostic screening of conventional rat holdings. The agent has occasionally been encountered in a range of other mammalian species (see reviews by Gaastra et al., 2009; Eisenberg et al., 2016a,b), and in the future it is likely additional strains will be delineated. The binomial name S. moniliformis, first suggested by Levaditi et al. (1925), will be used throughout this text in accordance with the ninth edition of Bergey’s Manual (Holt et al., 1994), although based on a relationship to the actinobacilli, Wilson and Miles (Collier et al., 1998) have used the name Actinobacillus moniliformis, which is here considered to be a synonym (for reviews, see Wullenweber, 1995; Eisenberg et al., 2016a,b). The foregoing citations (and Nelson, 1930a) describe the evolution of synonymy with this organism and the following list of names may be found there and in the older literature: Streptothrix muris ratti, Nocardia muris, Actinomyces muris ratti, Haverhilia multiformis, Actinomyces muris, Asterococcus muris, Proactinomyces muris, Haverhilia moniliformis, Actinobacillus muris, and Clostridium actinoides var. muris. Although a streptothrix-like organism had been recognized in the blood of some human patients with recurrent fever after a rat bite, the organism was not isolated in pure culture and characterized until 1914 (Schottmuller, 1914). Blake(1916) described the first etiologically confirmed case of human streptobacillosis in America. The association of the organism with human disease was clear from the beginning, but many years were to elapse before an understanding of the relationship was established between S. moniliformis and its natural reservoir host, the rat. The earliest associations of S. moniliformis and its rat host were in the context of respiratory disease, particularly pneumonia (Tunnicliff, 1916a,b; Jones, 1922; Turner, 1929), and in cases of otitis media in conventional rats (Nelson, 1930a,b; Koopman et al., 1991; Wullenweber et al., 1992). It was this respiratory context that was contributory to the confusion (discussed later in this chapter) between S. moniliformisLphase variants and M. pulmonis. Although Tunnicliff’s original isolates were from pneumonic tissues and, in addition, she was to claim that intraperitoneal injection of S. moniliformis caused pneumonias (Tunnicliff, 1916a,b), critical examination of her work casts doubt on whether the organism caused pneumonia or was merely reisolated from (preexisting) pneumonic tissues. Moreover, the weight of experimental and circumstantial evidence since has, to the contrary, established the inability of this organism to cause respiratory disease in rats (Nelson, 1930a; Strangeways, 1933; Boot et al., 1993, 1996).

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It was not until 1933 that Strangeways reported the asymptomatic occurrence of S. moniliformis in both wild and laboratory rats as a commensal of the nasopharynx. Streptobacillosis had inadvertently been caused in mice receiving Trypanosoma equiperduminfected blood sourced from the nasopharynx of rat donors. Subsequent investigations established the nasopharynx, oropharynx (and saliva) as the usual location of this organism in the rat and provided the ecological link to explain the injection of S. moniliformis into humans after a rat bite; at the time a puzzle of some significance. Swallowing of the organism by rat carriers results in contamination of the feces and urine with S. moniliformis and forms the basis for understanding how Haverhill fever (in humans) can result from milk or water contaminated with rat excreta (McEvoy et al., 1987; Bleich and Nicklas, 2008; Regnath et al., 2015). At the present time, S. moniliformis is believed to be a commensal with low potential for pathogenicity for the rat. It has not been etiologically associated with naturally occurring disease processes in this species except as a secondary invader. Experimentally, intravenous infections in rats have been used as a model of the infective arthritides (Lemer and Silverstein, 1957; Lemer and Sokoloff, 1959), but these bone and joint lesions have not been seen in the rat under natural circumstances. Although S. moniliformis occurred commonly in laboratory rats during the first half of the 20th century, variably as high as 50% (Nelson and Gowen, 1930; Strangeways, 1933; Bhandari, 1976), the incidence of S. moniliformis in the laboratory has markedly decreased in modern times under the influence of higher animal care standards and the general use of laboratory rats produced by gnotobiotic derivation. It continues, however, to be occasionally isolated (or its presence detected serologically or by PCR) from both wild (Boot et al., 1993; Easterbrook et al., 2008) and conventional laboratory rats (Matheson et al., 1955; Koopman et al., 1991) and, rarely, from hysterectomy-derived barriermaintained laboratory mice (Wullenweber et al., 1990a; Glastonbury et al., 1996). Rat-bite fever remains an occasional occupational hazard for laboratory personnel and the general public who have contact with pet or wild rats (Rake, 1936; Dawson and Hobby, 1939; Larson, 1941; Thjotta and Jonsen, 1947; Borgen and Gaustad, 1948; Hamburger and Knowles, 1953; Smith and Sampson, 1960; Holden and MacKay, 1964; Roughgarden, 1965; McGill et al., 1966; Gledhill, 1967; Gilbert et al., 1971; Anonymous, 1975; Biberstein, 1975; Anderson et al., 1983; McEvoy et al., 1987; Rumlely et al., 1987; Cunningham et al., 1998; Hagelskjaer et al., 1998; Downing et al., 2001; Mackey et al., 2014). The incidence of rat-bite fever deriving from laboratory rats is quite low. Of 65 cases summarized in one review (Elliott, 2007) only 8 (12%) were

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attributed to laboratory rat exposures. The last recorded case of human rat-bite fever acquired from laboratory rats occurred in 1983 (Anderson et al., 1983; Wullenweber, 1995; Eisenberg et al., 2016a,b). Laboratory rats may act as zoonotic reservoirs for S. moniliformis infections in laboratory mice (Freundt, 1959; Glastonbury et al., 1996) and in nature as reservoirs for wild mouse infections (Taylor et al., 1994). Unlike rat infections, mouse infections with S. moniliformis ordinarily are not lesionless and asymptomatic. Heritable variability in expression of disease has been observed with different inbred mouse strains (Wullenweber et al., 1990a), and more recently in black house rats (R.rattus) (Michel et al., 2018),. The mouse is not thought to be a carrier of commensal streptobacilli, and the routes of infection for mice are quite similar to those for humans. Infections in mice may be acute with generalized septicemia and lymphadenitis (erythema multiforme) or may be more chronic and characterized by osteomyelitis and polyarthritis, especially of the lower hind extremity and tail vertebrae (Mackie et al., 1933; Van Rooyen, 1936; Freundt, 1959; Kaspareit-Rittinghausen et al., 1990; Savage et al., 1984). Natural infection of pregnant mice may also result in abortions (Mackie et al., 1933; Sawicki et al., 1962). The mode of transmission from rat to mouse has been investigated but remains uncertain (Levaditi et al., 1932; Van Rooyen, 1936). Infections do not persist in mouse colonies rigidly separated from rats (Gledhill, 1967). In the mouse, as in humans, septicemia and polyarthritis may follow rat bites (Levaditi et al., 1932) or may be unrelated to this mode of transmission (Mackie et al., 1933; Van Rooyen, 1936). Similarly, Haverhill fever in humans (erythema arthriticum epidemicum) occurred as epidemics in Haverhill, Massachusetts, in 1926 and in Chester, Pennsylvania, in 1925, in which the source of infection was presumptively oral and eventually traced to infected milk to which rats had access (Parker and Hudson, 1926; Place et al., 1926; Place and Sutton, 1934; Editorial, 1939). Even in more recent times, human S. moniliformis infections may be unrelated to rat bites (Hazard and Goodkind, 1932; Sprecher and Copeland, 1947; Osimani et al., 1972; Lambe et al., 1973; McEvoy et al., 1987; Rumlely et al., 1987), although direct inoculation by rat bite or through skin abrasions appears to be the most common mode of transmission to humans. There are very few human cases on record of streptobacillary rat-bite fever after a mouse bite (Arkless, 1970; Gilbert et al., 1971). Human-to-human transmission has not been reported (Gaastra et al., 2009). Cultural isolation of S. moniliformis from clinical material is difficult, especially from nonlesioned rats, even when known to be positive serologically or by PCR (Gaastra et al., 2009; Boot et al., 2010). Especially with lightly infected carriers, cultural retrieval is difficult and

likely to result in false negative (Koopman et al., 1991; Boot et al., 1996). Recovery is impeded by the use of sodium polyanetholsulfonate (“liquoid”) in blood culture bottles and by overgrowth of biomic bacteria present in clinical swabs. Morphology of the organism is variable and pleomorphic, depending on environmental factors and age of the culture (Nelson, 1931; Heilman, 1941a,b). Smears from blood or other clinical materials most commonly show small (less than 1 mm wide and 1e5 mm long) Gram-negative rods and filaments (Francis, 1932).After subculture on 20% horse serum blood or infusion agar, slender interwoven masses of filaments 0.4e0.6 mm develop. Occasional filaments show spherical, oval, fusiform, or club-shaped swellings that may occur terminally, subterminally, or irregularly (hence the name “moniliformis”) by 6e12 h after subculture (Collier et al., 1998). After 12e18 h, the filaments fragment into chains of bacillary, coccoid, or fusiform bodies. Some 24e30 h after subculture on agar, tiny coccoid or ring forms are seen as well, accompanied by masses of bubbles and irregularly shaped cholesterol-containing droplets. Gram negativity is variable, and staining is more intense in the monilia-like swellings than in the filaments. In 20% horse serum infusion broth cultures, the morphology is similar, but aggregate filamentous masses are characteristically seen, giving the appearance of a coarsely granular sediment. The supernatant is typically clear, without turbidity, and contains small flocculent masses of organisms, giving the tube a snowflake or cotton ball appearance when the tube is agitated. Although described as a facultative anaerobe, ordinary aerobic incubator conditions suffice for isolation and propagation. Growth may be enhanced in 8%e10% carbon dioxide (CO2) environments, which are recommended (Smith and Sampson, 1960; Hansen, 2000). Colonies on serum or ascitic fluid infusion agar are 1e3 mm in diameter after 48e72 h of growth, circular, and low convex; may vary from colorless to grayish white; be translucent or opaque; and have a smooth, glistening appearance with butyrous consistency. Best results have been obtained by using trypticase soy agar with 20% horse serum (Smith and Sampson, 1960). On 5% sheep blood agar, colonies are barely visible at 24 h, are 1e3 mm by 48e72 h, and are smooth, gray, translucent, and essentially nonhemolytic. Gram-stained preparations of the growth show the characteristic morphology of the bacterium. Cystine trypticase broth, 10% horse serum, and 1% of various carbohydrates may be used for study of fermentative reactions (Smith and Sampson, 1960; Hansen, 2000). The organism is viable in serum broth and agar cultures for 2e4 days at 37
C, 7e10 days at 3e4
C, 14e15 days at 3e4
C in sealed cultures, and for several years in infected tissues, body fluids, or cultures maintained at from 25 to 70
C (Holt et al., 1994). Extensive phenotypic, cultural, and

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biochemical characterization of S. moniliformis isolates has been reported (Eisenberg et al., 2015a,b). It is believed that all isolates of S. moniliformis may undergo reversible conversion to transitional-phase variants. The term L1 was first used by Klieneberger (1935, 1936, 1942) to designate the occurrence in S. moniliformis cultures of small, filterable (Berkfeld V) variants morphologically resembling the bovine pleuropneumonia organism. Indeed, a sensation was caused when Klieneberger affirmed that the L1 variant of S. moniliformis belonged to the pleuropneumonia-like organisms (PPLOs). In this designation the “L” (for Lister Institute) was later applied to any bacterial variant reproducing in the form of very small cells lacking rigid cell walls. Individual isolates of S. moniliformis vary in degree of stability, and the L phase is typically unstable. L-phase variants differ from the bacillus in cellular and colonial morphology and in susceptibility to antibiotics, for example, penicillin, that act primarily on the cell wall. Occasionally, stable L variants may be achieved by manipulation, which do not undergo reversion to the parental type. Bacterial L variants may not be distinguished from true Mycoplasma colonies on the basis of size, “fried egg” appearance, or deep brown color in the central area (Freundt, 1959; McGee and Wittler, 1969). However, the usual reversion of L variants to parental forms on supportive media without inhibitors will ordinarily serve to identify bacteria (and is a useful proof to sustain diagnostic isolation of Mycoplasma). The L-phase variant with an incomplete cell wall is resistant to penicillin concentrations as much as 10,000 times higher than is the normal bacillus with cell wall (Razin and Boschwitz, 1968; McGee and Wittler, 1969), and is ordinarily only encountered in the laboratory after subculture, although it has been isolated directly from pneumonic rats (Klieneberger, 1938) and from the blood of penicillin-treated rat-bite fever patients (Dolman et al., 1951). When inoculated into mice, the L-phase variant is nonpathogenic; however, it frequently reverts in vivo to the bacillary form with full recovery of the pathogenic properties of the original S. moniliformis isolate. Great confusion was inadvertently introduced into the emerging understanding of the etiology of respiratory disease of laboratory rats when Klieneberger and Steabben (1937, 1940) isolated from pneumonic rats hitherto undescribed organisms that they termed L3 and believed to be L-phase variants of S. moniliformis. Similarly in 1937, Nelson described “coccobacilliform bodies” isolated from mice with respiratory disease, and several years later from rats (Nelson, 1940). Accumulated findings in these and other laboratories led to the conclusion that the L3 and Nelson’s coccobacilliform bodies were identical, and they have since been classified within the Mycoplasmatales as M. pulmonis

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(Freundt, 1957). It is now generally understood that L1 and L3 are not associated, that the L1 variant (and other L variants) of S. moniliformis are unrelated to the mycoplasmas (PPLOs), and that S. moniliformis and/or its L variants play no necessary role in murine respiratory mycoplasmosis. Opsonins, agglutinins, and complement-fixing antibodies have been demonstrated in the postinfective sera of both experimentally and naturally infected rats (Tunnicliff, 1916b), naturally and experimentally infected mice (Nelson, 1933; Van Rooyen, 1936; Merrikin and Terry, 1972), and human S. moniliformis infections (Brown and Nunemaker, 1942; Hamburger and Knowles, 1953; Robinson, 1963; Gilbert et al., 1971; Lambe et al., 1973). Use of Tween 80 in media (1.5%) used to propagate S. moniliformis for agglutination antigen appears to prevent the tendency of the organism to clump (Lambe et al., 1973). Agglutinins may not appear in promptly treated human cases (Holden and MacKay, 1964). Both direct (Lambe et al., 1973) and indirect (Holmgren and Tunevall, 1970; Lambe et al., 1973) immunofluorescence have been used to confirm S. moniliformis isolates and antibodies from rat-bite fever patients and laboratory rats (Lambe et al., 1973; Wullenweber et al., 1990b, 1992). The L-phase variant shares at least one common antigen with the bacillus, but it lacks at least one antigen present in the bacterial form (Klieneberger, 1942). An enzyme-linked immunosorbent assay (ELISA) procedure has been developed and is widely in use as a screening test to detect the presence of S. moniliformis in both rat and mouse colonies (Boot et al., 1993). Sufficient evidence is not available as to whether or not antibodies invariably occur in asymptomatic carrier rats to make serological screening reliably useful in monitoring laboratory rat colonies (i.e., validity of serologic negative), but conventional rats were almost invariably found to be ELISA positive (Boot et al., 1993). A compounding immunodiagnostic issue is that rat strains are known to vary in intensity of antibody responses to infection (Boot et al., 2010). An immunoblot (Western blot) procedure has been developed to confirm antibodies detected by ELISA. The antigenic relatedness of S. moniliformis isolates from various species has been investigated to some extent. Isolates of S. moniliformis from mouse, rat, and human infections were cross-reactive by ELISA (Boot et al., 1993). Varying degrees of electrophoretic protein patterns were shown among human, rodent, and avian isolates (Costas and Owen, 1987). Such isolates may have varying biological characteristics. Guinea pig isolates formerly attributed to S. moniliformis (Smith, 1941; Aldred et al., 1974) have more recently been determined by analytic PCR to be due to cross-reactivity with the guinea pig agent Leptotrichia spp. (Boot et al., 2007).

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Doubt exists as to whether guinea pig isolates are truly S. moniliformis or, rather, taxonomically removed. Bergey’s Manual classified such organisms as Sphaerophorus caviae (Kirchner et al., 1992; Holt et al., 1994). Diagnosis is established by isolation and cultural characterization of S. moniliformis. Important confirmatory information may be obtained by PCR with swab samples derived from multiple clinical sites, for example, pharynx, trachea, and lymph nodes (Boot et al., 2002). For a number of reasons, prominent among them the stated difficulty of cultural retrieval, PCR has increasingly gained favor as the principal means of detection of this organism in both human and rodent clinical material (Boot et al., 2002; Elliott, 2007; Gaastra et al., 2009). PCR may be used to screen rodent colonies for the presence of S. moniliformis and has been used in surveys of wild rat microbiology (Kimura et al., 2008). PCR is widely used to diagnose rat-bite fever in human clinical material (Berger et al., 2001; Wallet et al., 2003; Mignard et al., 2007; Nakagomi et al., 2008; Mackey et al., 2014). Primer sets have been developed to identify S. moniliformis down to genus level for screening purposes (Berger et al., 2001; Boot et al., 2002; Wallet et al., 2003) but speciation requires more detailed and systemic confirmatory molecular analysis and has been facilitated by the development of MALD-TOF MS (Eisenberg et al., 2015a,b, 2016a,b). Older diagnostic strategies also used intraperitoneal and foot pad inoculation of test mice with causation of septicemia and embolically distributed lesions, including polyarthritis from which S. moniliformis could be reisolated. With regard to treatment, little is known about the efficacy of antibiotic treatments in abolishing latent infections of the nasopharynx in rats, nor has this strategy been of interest. Treatment of colonies with important zoonotic infection should be discouraged. Considerable information is available, of course, with regard to antibiotic treatment of S. moniliformis infections in humans (Roughgarden, 1965; Lambe et al., 1973; Wullenweber, 1995; Cunningham et al., 1998; Downing et al., 2001; Elliott, 2007; Gaastra et al., 2009). However, information about systemic treatment of embolic foci (in humans) is of little use in predicting treatment methods for the rat, in which the ecology of the organism is quite different. There is no evidence that the organism may hematogenously traverse the rat uterus or reside as a commensal in the rat reproductive tract. Infections have not been reported to occur in rats derived by gnotobiotic means and shielded from proximity to conventional rats.

B. Spirillum minus (Spirochetosis) The status of S. minus as a bacterial pathogen of rats, as with S. moniliformis, is somewhat uncertain. It is carried

asymptomatically by rat and mouse hosts. Similar to S. moniliformis, the nonpathogenicity of S. minus for its natural hosts has been overshadowed by the interest attending this organism as a cause of rat-bite fever in humans. Although illness after rat bites had been known since ancient times in India, where it is believed to have originated (Row, 1918), and had been known for centuries in Japan as sodoku (so meaning rat; doku meaning poisoning), it was not until 1902 that the disease was described by Japanese workers in European journals (Miyake, 1902) and given the name “Rattenbisskrankheit,” or rat-bite fever. The article by Miyake (1902) led to the recognition of rat-bite fever as a clinical entity and to the realization that a number of previous accounts of “poisoning” after rat bites had been made from America and the European continent (for early reviews, see McDermott, 1928; Robertson, 1924, 1930). Before Miyake’s report, it had been recognized that wild Ratti norvegicus (Mus decumanus) could carry spirillar organisms in the blood (Carter, 1888), and slightly later in mice (Breini and Kinghorn, 1906; Wenyon, 1906). The association was left, however, until 1915, when Futaki and his coworkers discovered this cause of rat-bite fever in humans (Futaki et al., 1916). Readers will recall that only 1 year before, S. moniliformis also had been associated with rat-bite fever in humans (Schottmuller, 1914), and a controversy was initiated that continued for almost three decades as to the “true” cause of rat-bite fever. As so often happens in science, the passage of time and the careful documentation of clinical cases and laboratory data were to reveal that both of these organisms would be recognized as causing similar, but nonetheless different, syndromes of “fever” after rat bites (Brown and Nunemaker, 1942). The evidence was summarized at about the same time by Allbritten et al. (1940) and Farrell et al. (1939), who carefully distinguished three diseases: sodoku and streptobacillosis after rat bite, and Haverhill fever after (presumed) ingestion of S. moniliformis (for reviews, see Adams et al., 1972; Watkins, 1946;Gaastra et al., 2009). Spirillary sodoku and streptobacillosis are clinically differentiated on the basis of (1) longer incubation period (14 days); (2) formation of an indurated “chancre,” or ulcer, with regional lymphadenitis and lymphangitis deriving from the bite site; (3) more regularly periodic recurrent fever; (4) maculopapular, erythematous rash spreading from the initial lesion (Downing et al., 2001); (5) the absence or rare occurrence of petechiae; (6) the absence or rare occurrence of polyarthritis in the case of sodoku (S. minus infection); and (7) lower risk of mortality (6.5%) than streptobacillary rat-bite fever. Streptobacillosis, on the other hand, was usually shorter in onset (within 10 days) and septicemic, with morbilliform or petechial hemorrhages, endocarditis, irregularly recurrent fever, and polyarthritis. All

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known cases of rat-bite fever contracted from laboratory rats and mice have been of the streptobacillary type (Gledhill, 1967; Gaastra et al., 2009). Human rat-bite fever of the S. minus type, in India and Brazil, has a seasonal epidemiology (Chopra et al., 1939; Cole et al., 1969; Bhatt and Mirza., 1992). The organism has been identified in a number of other hosts, including dog, cat, weasel, and ferret, although it is believed that carnivore infections represent zoonoses contracted from rodent carriers in the course of predation (Hata, 1912). At present only one species, S. minus, is taxonomically recognized (Holt et al., 1994). The older literature, however, contains references to Spirochaeta morsus muris, Spirillum minor, Spirochaeta laverani, Spironema minor, Leptospira morsus minor, Spirochaeta muris, and Spirochaeta petit, which are all regarded as synonyms. The frequent early references to this organism as a spirochete were based not only on morphologic criteria: indeed, it is a matter of some historic interest that Ehrlich used the mouseeS. minus system in experiments leading to the development of salvarsan for the chemotherapy of human syphilis (McDermott, 1928), that early cases of S. minus rat-bite fever in humans were successfully treated with salvarsan (Hata, 1912; Surveyor, 1913; Dalal, 1914; Spaar, 1923), and that the organism has been advocated as a model system for the study of syphilitic infections (Stuhmer, 1929). S. minus is classified by Bergey’s Manual within the Spirillaceae, and it appears to be the only species in the genus that is both pathogenic and, at present, uncultivable on artificial media (Holt et al., 1994; Gaastra et al., 2009). For that reason, isolation of S. minus requires inoculation of test animals (mice, rats, or guinea pigs). As identified in fixed and stained blood smears from infected humans or laboratory rodents, the cells are Gram negative, appear short and thick but are approximately 0.5 mm in width and 1.7e5 mm in length, and have two to six spirals (for illustrations, see BayneJones, 1931). The organism has tufts of flagella (MacNeal, 1907; Adachi, 1921) at each end and is actively motile. The little that is known about the pathogenicity of S. minus in its natural hosts (rats and mice) is related to the diagnostic inoculation of laboratory rats and mice with blood from rat-bite fever patients (or strains of S. minus initially derived from such patients). Although some surveys of both wild and laboratory rats and mice have indicated the prevalence of the organism in the decades surrounding World Wars I and II (de Araujo, 1931; Francis, 1936; Knowles et al., 1936; Brown and Nunemaker, 1942; Jellison et al., 1949; Humphreys et al., 1950; Dolman et al., 1951), these surveys have not reported the occurrence of lesions referable to S. minus. The presence of unrecognized intercurrent

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disease in experimental laboratory hosts was so prevalent at the time these studies were, the interpretation of clinical signs and lesions said to be caused by S. minus must be viewed with circumspection. Little is known about the epidemiology of this organism. In modern times, it still occurs as an infrequent cause of wild rat-associated rat-bite fever, predominantly in Asia, but based on case reports, distributed globally. The experimental disease has been studied by the intraperitoneal injection of blood into laboratory rats, mice, guinea pigs, rabbits, cats, ferrets, and rhesus monkeys. These studies achieve a fair consensus on placing the incubation period in experimental hosts at about 3e5 days or longer (Wenyon, 1906; Dalal, 1914; Leadingham, 1928, 1938; McDermott, 1928; Stuhmer, 1929). The height of infection in terms of demonstrable organisms by dark-field examination of thick films occurs 2e 3 weeks after inoculation of experimental rats, mice, and guinea pigs. Experimental animals may remain infective 6e12 months after inoculation (Leadingham, 1938). Few, if any, signs other than an initial bacteremia have been reported in mouse hosts (MacNeal, 1907; McDermott, 1928). In the rat and guinea pig, however, subcutaneous S. minus infection may typically also cause indurated or ulcered chancroid lesions at the inoculation site, with regional lymphadenopathy and fever reminiscent of human sodoku-type rat-bite fever (McDermott, 1928). Spirochetes frequently are demonstrable in the serous discharge from primary lesions (McDermott, 1928). In considering the essential features of the experimental disease in rats, McDermott (1928) described the following successive phases: incubation, development of primary inflammatory lesion, and lymphadenopathy with organisms initially limited to these tissues; a secondary septicemic stage with organisms demonstrable in the blood; a latent stage in which the blood was free of (demonstrable) spirochetes and there were no obvious lesions; and, finally after many months, a tertiary stage with gummatoid lesions (abscesses and granulomas) of the lungs, spleen lymph nodes, and liver. Gummas have been described as long-term lesions by others as well (Stuhmer, 1929; Leadingham, 1938). The gummatoid lesions particularly are suspected of being otherwise induced and merit reexamination. It is worth restating, however, that none of these lesions has been reported as occurring in naturally infected rat hosts. Diagnosis of S. minus infection may be made by demonstration of the actively motile organisms in wet mounts made of primary lesion exudates or peripheral blood by dark-field or phase-contrast microscopy. Blood films stained with Wright’s, Giemsa, or silver impregnation stains also should be examined. Because it is recognized that septicemic phases may be brief, failure to demonstrate the organism in peripheral blood should always be supported by culture (to isolate S. moniliformis,

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if present) and animal inoculation. It is recommended that four mice and two guinea pigs be used to screen each suspect blood sample and that preinoculation blood samples be examined from the test animals to preclude use of intercurrently infected mice or rats (Brown and Nunemaker, 1942; Rogosa, 1974). Suspect blood should be inoculated subcutaneously, intraperitoneally, and intradermally (Wilson and Miles, 1975). Blood and peritoneal fluid from the test animals should be examined microscopically at weekly intervals for 4 weeks. Peritoneal fluid is usually richer in organisms than is blood (Jellison et al., 1949). An equal number of animals should be identically inoculated and examined by using an aliquot of suspect blood heat treated to 52
C for 1 h (Rogosa, 1974). The heart and tongue of test animals at the conclusion of the test period (4 weeks) should be used to make Giemsa-stained impression smears. Organisms are often present in fair numbers in tongue and cardiac tissue when scarce (as in latent phases) in blood or peritoneal fluid (Jellison et al., 1949; Rogosa, 1974; Wilson and Miles, 1975). Live organisms in fresh preparation have been immobilized by immune sera (McDermott, 1928), but serologic tests have not been advocated for diagnosis because of the technical problems encountered in making antigenic reagents. Approximately one-half of human cases are positive by Treponema pallidum serodiagnostic methods, and the importance of taking a serum sample early in the course for this purpose has been stressed (Woolley, 1936). Rabbits infected with S. minus blood suspensions have been reported to develop positive WeileFelix reactions with Proteus OXK strains (Savoor and Lewthwaite, 1941). The mechanism of zoonotic transfer of S. minus from infected rats via bite lesions is unknown. The organism has not been demonstrated in rat or mouse saliva, although this question has been investigated (McDermott, 1928; Leadingham, 1938). The organism is present in high numbers in tongue and myocardium of infected animals and salivary glands of infected mice (Bok R., 1940, cited by Gaastra et al., 2009). It has been observed in urine of human rat-bite fever cases (Leadingham, 1938), as well as isolated (by inoculation) from the urine of naturally and experimentally infected rats (Humphreys et al., 1950). The current hypothesis suggests that bleeding oral abrasions at the time of the bite may result in bite inoculation. There have been only two human cases on record after a mouse bite despite the documented widespread distribution of S. minus in both wild and conventional laboratory mouse populations (Jellison et al., 1949). The organism has not been reported in laboratory animals in modern times, and it is unknown therefore whether it can be vertically transmitted to gnotobiotically derived animals.

Chemotherapy of naturally acquired or experimental S. minus infection has not been reported for the rat in recent times, although human cases have been reported as successfully treated with penicillin or streptomycin (Jellison et al., 1949; Roughgarden, 1965). A battery of antibiotics has been assessed for efficacy in the prevention of transmission of S. minus from infected donor mice to treated mouse recipients (Tani and Takano, 1958).

C. The Streptococci The Genus Streptococcus is classified in the Phylum Firmicutes, Class Cocci, Order Lactobacillales, Family Streptococcaceae. The streptococci are facultatively anaerobic, catalase negative, Gram-positive cocci that occur in pairs or chains. While several of the species are pathogenic and responsible for serious animal and human infections, many are not, and in fact are commensal components of the microbiome of the naso- and oropharynx, the intestines, and skin. The streptococci may be identified by genomic 16s rRNA homology (Facklam, 2002). Dendrograms of these relationships do not necessarily correlate with phenotypic types of hemolysis or Lancefield groups as are outlined in the following. More commonly, identification of streptococcal species from clinical material proceeds from consideration of the phenotypic type of hemolysis by isolates on blood agar: 1. a-Hemolysis refers to the zone of darkish, greenish partial hemolysis produced by S. pneumoniae and by the viridans group of commensal streptococcal species (including Streptococcus mutans). 2. b-Hemolysis is defined by the zone of clear hemolysis surrounding colonies of this type caused by production of the exotoxin streptolysin. The b-hemolytic streptococci are further classified phenotypically based on capsular surface antigens: the Lancefield groups. These groups include group A, the Streptococcus pyogenes group, group B, the Streptococcus agalactiae group, group C, the Streptococcuszooepidemicus and Streptococcus equi groups, and groups F and G, the Streptococcus dysgalactiae group and group K. The former group N has been reclassified as a separate genus, the Lactococcus. 3. g-Hemolysis, by which is meant no, or only slight, hemolysis on blood agar. The g-hemolytic groups include the former Lancefield group D that is now recognized as a separate genus, the Enterococcus. Several of the streptococci important in terms of health of laboratory rats are listed among the bacterial agents recommended for rat health surveillance by FELASA and AALAS (Table 12.1).

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1. Streptococcus pneumoniae In conformity with the ninth edition of Bergey’s Manual (Holt et al., 1994), the name S. pneumoniae has been adopted for this organism, and the former names Diplococcus and Pneumococcus regarded as synonyms. In the Introduction to this chapter it was pointed out that S. pneumoniae was unrecognized as a pathogen of laboratory rats during the first half of the 20th century. The organism appears not to have been encountered even in microbial surveys of respiratory flora from laboratory rats during this period (Nelson, 1930a,b;Block and Baldock, 1937), unlike similar surveys undertaken later (Matheson et al., 1955; Weisbroth and Freimer, 1969). As will be described in more detail ahead, in the latter half of the 20th century, S. pneumoniae had been observed in the context of acute respiratory episodes in rats, most commonly, but not limited to, those in the adolescent and young adult age groups. The evidence does not suggest that these infections occurred and were unrecognized earlier, but rather that fundamental changes in the rat host occurred later, perhaps in the nasopharyngeal microbiome, rendering it more susceptible to this organism. It also needs to be recognized that S. pneumoniae is quite commonly carried as an asymptomatic but regulated pathobiont of the nasopharyngeal microbiome of humans coming in contact with laboratory rats (Lemon et al., 2010; Cremers et al., 2014; Piters et al.,2016; Chan et al., 2016). While pointed out later that several of the pneumococcal types have occurred asymptomatically in rats, there is little evidence, if any, that S. pneumoniae is a normative pathobiont of rats as it is in humans. The first report of respiratory epizootics in laboratory rats caused byS. pneumoniae was published in 1950 (Mirick et al., 1950). However, as late as 1963 it was written that acute respiratory episodes in rats of any age were rare (Tuffery and Innes, 1963), and as late as 1969 that bacterial pneumonias, including those caused by S. pneumoniae, were unlikely to be encountered in properly managed facilities (Brennan et al., 1969b). It was regarded as a widespread and major primary bacterial pathogen of laboratory rats 40e50 years ago (Weisbroth and Freimer, 1969), but the incidence of infection of laboratory rats by S. pneumoniae has been much reduced in more recent years, principally as a result of common availability of barrier-reared disease-free rat stocks. There have been no reports of clinical disease caused by S. pneumoniae in more than 40 years. The pathogenicity of certain serotypes of the organism is sufficient to serve as a primary cause of disease in the absence of physiologic stress or other infectious incitants, although it has been recognized as frequently in concurrent infection with Mycoplasma. Certain serotypes appear to cause essentially asymptomatic infection (Fallon et al., 1988).

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The organism was documented to be widespread among commercial rat breeding stocks in 1969 (Weisbroth and Freimer, 1969). In the latter study, 19 of 22 commercial sources were found to be offering for sale adolescent rat carriers of S. pneumoniae with varying degrees of clinical signs. The majority of animals from which pneumococci were isolated were asymptomatic carriers of the organism in the nasoturbinate mucosa. It was concluded that the organism is generally carried in the upper respiratory tract, particularly the nasoturbinates and middle ear in the absence of clinical signs. Of 20 rats without clinical signs, no pneumococci were isolated from pulmonary tissues, whereas S. pneumoniae was isolated from 17 in nasoturbinate washings. The infection is not ordinarily inapparent, however, and clinical signs of respiratory disease with varying degrees of mortality are more typical (Kelemen and Sargent, 1946; Innes et al., 1956; Ford, 1965; Baer, 1967; Baer and Preiser, 1969; Mitruka, 1971; Tucek, 1971; Adams et al., 1972). Serosanguinous to mucopurulent nasal exudates are frequently the first signs of clinical illness and ordinarily precede pulmonary involvement. Rhinitis, sinusitis, conjunctivitis, and otitis media are common gross lesions of upper respiratory infection. Outward bulging of the tympanic membrane may be noted as a sign of pus under pressure in the middle ear. Histologically, mucopurulent exudates are seen to overlay the respiratory mucosa of the turbinates, sinuses, eustachian tube, nasolacrimal duct, and tympanic cavity. Acute inflammatory cells, primarily neutrophils, and more chronic inflammatory cells, including plasma cells and lymphocytes, infiltrate the mucosa and underlying submucosa of these tissues. Abundant numbers of organisms may be observed in the exudates and superficial levels of the mucosa by means of tissue Gram stains or smears. Concomitant clinical signs include postural changes (e.g., hunching), abdominal breathing as pneumonia supervenes, dyspnea, conjunctival exudation, anorexia with loss of weight, depression, and gurgling or snuffling respiratory sounds (rales). The onset of clinical signs and lesions is more often acute or subchronic than chronic and affects rats of all ages but particularly younger age groups. The ordinary route of progression of upper respiratory infections is by descent from the nasopharynx to pulmonary tissues. Frequently, the right intermediate lobe of the lung is the first to be affected. Initially, fibrinous bronchopneumonias in certain lobes develop rapidly into classical confluent fibrinous lobar pneumonia. Frothy, serosanguinous fluid exudes from the cut trachea. Microscopically, the mucosa of the trachea, bronchi, and bronchioles is necrotic and eroded, with fluid and purulent exudates, often with blood, in the lumina. The alveolar capillaries are congested, and alveoli themselves are filled with blood, proteinaceous

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fluids, neutrophils, or variable combinations of these. Tissue Gram stains reveal abundant Gram-positive cocci in affected tissues, in microcolonies, as freely distributed single cells, and intracellularly within phagocytic cells, including neutrophils. Death may supervene at this point depending on the degree of pulmonary involvement. Frequently, however, the organism escapes to the thoracic cavity, and lesions of fibrinous pleuritis, pleural effusion, and fibrinous pericarditis are common. Organisms may ascend from the thoracic organs to become septicemic, and septicemia is a frequent terminating event. Less commonly, the organism is embolically distributed to any of a variety of organs and locations, and such complications as purulent arthritis; focal necrosis and/or infarction of the liver, spleen and kidneys; and fibrinopurulent peritonitis, orchitis, and meningitis have been observed (Baer and Preiser, 1969; Weisbroth and Freimer, 1969; Mitruka, 1971). It is known that the serum biochemistry of naturally infected rats is disturbed with increases in several serum enzymes (including glutamic-pyruvic transaminase, glutamicoxaloacetictransaminase, and lactate dehydrogenase) and a- and b-globulins (Mitruka, 1971). Diagnosis is established by retrieval of S. pneumoniae from affected tissues and is supported by characteristic gross and microscopic lesions. Rats may be screened for S. pneumoniae by the culture of nasopharyngeal swabs (Mirick et al., 1950; Accreditation Microbiological Advisory Committee Accreditation, 1971) or saline washings of the tympanic cavity and nasoturbinates (Weisbroth and Freimer, 1969), because these upper respiratory locations in carrier rats are more likely to yield the organism than are pulmonary tissues. Both quantitative nasopharyngeal cultures and qPCR (compared with nasopharyngeal swabs) have been established as yielding somewhat higher numbers of positive rat carriers, but neither of the former quantitative methods was compared with nasopharyngeal wash cultures (Kontiokari et al., 2000). Primary clinical samples (swabs, washings, exudates, heart blood, etc.) are plated directly onto 5% blood agar. Growth is facilitated by 10% CO2, and some isolates require microaerophilic environments (Wilson and Miles, 1975). Colonies of capsulated pneumococci on blood agar are raised, circular, and 1e2 mm in diameter with steeply shelving sides and an entire edge. Early in incubation, the colonies are domeshaped and glistening (especially Type 3), but by 24e48 h, the centers collapse because of autolysis, giving a typical concave umbilication to the top of the colony. S. pneumoniae colonies are surrounded by a small (alpha) zone of hemolysis with greenish (viridans) discoloration in the media. The morphology of S. pneumoniae is consistent both from clinical materialsdfor example, smears of exudates or tissuesdand from subcultivation on artificial

media. The individual cells are ovoid or lanceolate and typically occur as pairs, with occasional short chains, surrounded by polysaccharide capsules. The capsule may be demonstrated by Gin’s method (Baer and Preiser, 1969). Gram-staining reactions may be inconsistent from clinical materials; however, the majority of smears and isolates are distinctly Gram positive with clear nonstaining capsular material surrounding the cells and are of value in presumptive identification. Suspicious colonies from primary isolates should be individually picked and established in isolation by subculture on blood agar. Identity of the isolate should be confirmed by characteristic reactions in differential media, sensitivity to optochin and bile solubility, and serological typing (Nakagawa and Weyant, 1994). The API 20 Strep kit and similar identification systems can be used for identification of primary isolates (Hansen, 2000). Optochin (hydrocuprein hydrochloride) is used to distinguish S. pneumoniae (optochin sensitive) from the other a-hemolytic viridans streptococci (noninhibited). Sensitivity is established by a zone of inhibited growth surrounding optochin (5 mg)-impregnated paper discs placed on the surface of subcultures evenly streaked on blood agar plates (Bowers and Jeffries, 1955). Sensitive isolates are inhibited for a zone of about 5 mm from the edge of the disc. Bile solubility is another useful test to confirm isolation of S. pneumoniae. Isolates that are bile soluble will clear (i.e., autolyze) during this period, and it is generally understood that bile solubility simply acts to accelerate the natural autolytic process that causes central collapse of colonies on blood agar (Wilson and Miles, 1975). Bile solubility of S. pneumoniae isolates is more variable than is optochin sensitivity (Accreditation Microbiological Advisory Committee Accreditation, 1971; Wilson and Miles, 1975). More recently use of the MALDI-TOF MS procedure has come into play as a method to rapidly identify isolated S. pneumoniae and to differentiate the isolate from other S. mitis group streptococci (Harju et al., 2017; Werno et al., 2012). Biochemically confirmed isolates of S. pneumoniae may be typed by the use of type-specific antisera in the NeufeldeQuellung reaction (Austrian, 1974), although serotyping is not required for the diagnosis of S. pneumoniae infection. Quellung reactions are based on swelling of the capsules surrounding individual organisms in the presence of specific antisera and may be performed by the microscopic examination of cover-slipped slides with a mixture of one loopful each of an overnight ToddeHewitt broth culture, specific antiserum, source of the infection. Weisbroth and Freimer (1969) found that rat isolates from various colonies were usually monotypic and that the same type appeared to prevail in distinct geographical patterns. Most commonly, types 2, 3, and 19 (Mirick et al., 1950;

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Innes et al., 1956; Baer and Preiser, 1969; Weisbroth and Freimer, 1969; Mitruka, 1971; Adams et al., 1972)dand less commonly types 8 and 16 (Ford, 1965; Baer, 1967)d have been clinically encountered in rats on the American continent. The pneumococcal type may also bear on pathogenicity for the rat. Type 35, for example, has been documented as being carried in the nasopharynx by rats (and even mice) as an inapparent agent in the absence of disease (Fallon et al., 1988). Virulence of S. pneumoniae is known to be highly correlated with capsular material (Nakagawa and Weyant, 1994). Although it is recognized that rats respond with antibodies to natural infections with S. pneumoniae, this property has not received a great deal of attention as a diagnostic method, perhaps because of the ease with which the organism may be culturally retrieved. Several serologic tests have been reported (Lund and Henrichsen, 1978; Schiffman et al., 1980; Jalonen et al., 1989), but none have been recommended for use in health surveillance of laboratory rats. It has been established that certain rat stocks are more resistant to certain pneumococcal types and that young animals are more susceptible than are mature rats (Ross, 1934). Natural resistance occurs to experimental infection with pneumococcal types commonly found in rats, e.g., types 2 and 3 (Ross, 1931, 1934). As with serologic tests, although both direct and indirect immunofluorescence have been used to locate S. pneumoniae in infected tissues (Coons et al., 1942; Kaplan et al., 1950), and pneumococcal antigens have been detected by counterelectrophoresis of clinical specimens (Spencer and Savage, 1976), these methods have not received attention as routine diagnostic methods in rats. Laboratory mice are very sensitive to the intraperitoneal injection of clinical samples or broth cultures containing S. pneumoniae (Rake, 1936; Morch, 1947; Cowan, 1974; Lennette et al., 1974), with death often occurring in 24e72 h with dilutions containing as few as five microorganisms (Ford, 1965). In the past, mouse inoculation had been advocated as a biological sieve for human diagnostic specimens, especially where speed was of the essence (Rathbun and Govani, 1967), but it has not been advocated as a method to diagnose rat infections (Accreditation Microbiological Advisory Committee Accreditation, 1971; Nakagawa and Weyant, 1994). Antibiograms of isolates of S. pneumoniae from the rat have generally established the susceptibility of the organism to antibiotics effective against streptococci (Osimani et al., 1972). Antibiotic sensitivity should be established before treatment, because penicillinresistant isolates have been retrieved from naturally infected rats (Mirick et al., 1950). Most isolates are sensitive to penicillin, however, and benzathine-based penicillin has been used for effected treatments in the

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course of epizootics (Baer, 1967; Weisbroth and Freimer, 1969; Adams et al., 1972). Rats have also been effectively treated by intraperitoneal injection of type-specific (rabbit) antiserum (Mirick et al., 1950). It has been generally agreed that antibiotics are useful in the amelioration or prevention of systemic infection (septicemia, pneumonia); however, antibiotics are thought to be ineffective in eradication of organisms from infected rat colonies, because of rebound after withdrawal of treatment. It was concluded (Weisbroth and Freimer, 1969) that the bacteria in and on the mucosa of the upper respiratory tract are shielded from antibiotic access. The organisms may be excluded by establishment of barrier colonies of gnotobiotically derived rats. No pneumococci were isolated from commercial sources of rats thus derived in the course of a microbiologic survey (Weisbroth and Freimer, 1969). Because of the importance of S. pneumoniae in human health, rats have been used extensively as the host in model infection systems (see review by Chiavolini et al., 2008). Such studies have established a number of factors bearing on susceptibility, including splenectomy (Biggar et al., 1972; Boart et al., 1972; Leung et al., 1972), age (Ross, 1931, 1934), genetic stock (Ross, 1931), iron deficiency (Shu-heh et al., 1976), and pulmonary edema (Johanson et al., 1974). Other reports have explored the effect of S. pneumoniae infections on such host systems as nitrogen metabolism and protein synthesis (Powanda et al., 1972), serum proteins and enzymes (Mitruka, 1971), blood pH and electrolytes (Elwell et al., 1975), hepatic enzymes (De Rubertis and Woeber, 1972; Canonico et al., 1975), thyroid physiology (Shambaugh and Beisel, 1966), and the effect of antihyperlipidemic agents, e.g., clofibrate (Powanda and Canonico, 1976). 2. The b-Hemolytic Streptococci As mentioned earlier, the b-hemolytic streptococci are typed by Lancefield antisera into groups A, B, C, F, G, and K. While the group A streptococci of the S. pyogenes group have in the past been observed as a pathogen of laboratory mice (cited by Geistfeld et al., 1998) there seem to be no records of this group infecting laboratory rats. Similarly, streptococci of group C, the S. zooepidemicus group, have often been associated with pathogenicity in guinea pigs, but there appear to be no reports of rat infection by this group. The group B streptococci (GBS) of the S. agalactiae group on the one hand are generally regarded as commensals of the human genitourinary, oropharyngeal, and intestinal microbiome; on the other hand, they can cause serious human and animal diseases. Streptococcal b-hemolytic isolates presumptively identified as group B based on latex agglutination with Lancefield typing antisera can be confirmed as group B based on being CAMP test positive and also by failure to hydrolyze

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hippurate. The CAMP test is based on observation of enhanced hemolysis in synergy with staphylococcal b-hemolysin when grown on blood agar. The GBS were first identified as causative agents of bovine mastitis and the CAMP test was developed to presumptively confirm field isolates. As mentionedearlier, the agent is carried as a commensal in, variably, up to 40% of human genitourinary tracts. GBS is recognized as an important cause of severe neonatal human infection during the first week of life (early onset) or somewhat later, i.e., from 1 week to 3 months of age (late onset) (Onile, 1985). While encountered more commonly as septicemia in laboratory mice (Geistfeld et al., 1998), GBS have been less frequently observed as septicemic infections of both neonatal (Shuster et al., 2013) and adult female laboratory rats (Bodi Winn et al., 2017). In the latter report, GBS were isolated from the nasopharynx of asymptomatic adult males as well. Group G streptococci (GGS) isolates are presumptively differentiated from the GBS by negative CAMP test and confirmed as group G by Lancefield typing antisera. The type species is S. dysgalactiae (Woo et al., 2001) with a number of subspecies. The subspecies Streptococcus equisimilis and Streptococcusmilleri occur as commensals of the human microbiome, while the subspecies Streptococcuscani and Streptococcusintestinalis have as reservoirs microbiomes of the dog and swine, respectively. There appears to be a common factor of inductive stress preceding clinical episodes of GGS infections of laboratory rats. GGS seemingly carried as oropharyngeal commensals were observed to have embolically caused endocarditis following dental extraction in rats with periodontal disease (Morellion et al., 1988). In the other reported episode, adolescent rats developed lymphadenitis 10 days following gavage with 7,12-dimethylbenz(a)anthracene (Corning et al., 1991). 3. Enterococcus sp. Infection Although traditionally regarded as a streptococcal genus, the fecal streptococci have been classified as a separate genus, Enterococcus. The enterococci are catalasenegative and facultatively anaerophilic, form pinpoint colonies, usually exhibit slight a-hemolysis, and occur in chains. Isolates are serologically (weakly) typed as Lancefield group D and fall into two groups: those able and those unable to grow on tellurite-containing media. Because the common species occurring in rodents will not grow on tellurite-containing media, the latter should not be used for isolation of Enterococcus. Primary isolation may be easily accomplished by incubation of fecal samples (or swabs) directly onto eosin methylene blue agar. Suspicious colonies can be picked for isolation and propagated in pure culture in trypticase soy broth (Hoover et al., 1985). The enterococci may be speciated by commercial kits (e.g., API a-Strep or similar).

Rodent clinical infections have mainly been identified as occurring within the phenotypic group III of enterococcal species closely related to Enterococcus faecium, including Enterococcus durans, Enterococcus dispar, and Enterococcus hirae (Gades et al., 1999; Teixiera et al., 2001). This group is unable to grow on telluritecontaining media and is usually negative for acid on mannitol, sorbitol, and sorbose. Subsequent work with new and referred isolates of Enterococcus from rats have established a new species, Enterococcus ratti (originally named Enterococcus rattus by Uhl et al., 1998), closely related to E. durans and E. hirae as the common cause of naturally occurring neonatal diarrhea in rats (Teixiera et al., 2001). Phenotypic and genomic characteristics of these group III strains, enabling cultural differentiation, SDS-PAGE dendrography, and studies of DNAeDNA relatedness, have been reported (Teixiera et al., 2001). As also observed by Etheridge and Vonderfecht (1992), E. ratti isolates are usually slow growing with weak or delayed reactions in culture (Teixiera et al., 2001). Enterococci are ubiquitous in nature and commonly carried as bacterial commensals in the human gastrointestinal tract. They are of increasing importance as causes of nosocomial hospital infections of patients on antimicrobial regimens. The most common species in humans, E. faecium and Enterococcus faecalis, are likewise most commonly carried by laboratory rats (and the origins of contamination by Enterococcus in rederived rats removed from isolators, as with S. pneumoniae, are thereby suggested). Most clinical episodes have been reported as occurring in accreditable animal care environments in rats known to be free of common pathogens (Table 12.1). In rats also, enterococci are ordinarily carried in the gastrointestinal tract as nonpathogenic commensals and may even be regarded as having a probiotic function in that their abundance competes with other forms to act as an inhibitor on ascendancy of other bacterial pathogens (Hansen, 2000). In a number of instances, enterococci have been diagnosed as causal in an enteropathic syndrome of diarrhea in neonatal rats. The indicted species include E. durans (formerly S. faecium durans-2) (Hoover et al., 1985), as well as Enterococcus-like agents not further identified (Etheridge and Vonderfecht, 1992; Gades et al., 1999), but as outlined earlier, they are now all regarded as probable E. ratti infections (Teixiera et al., 2001). Clinical episodes have tended to occur in the 6- to 12-day age group with high morbidity and low mortality. Clinical signs have typically included abdominal distension and rough, yellowish matted haircoats, with perineal scalding and soiling, and ulceration of the skin of the hind limbs (Gades et al., 1999). The cages of affected litters have wet bedding and are frequently devoid of formed feces. Dams with affected litters may also have soft feces (Hoover et al., 1985), and in the case reported

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by Gades et al. (1999), the dam was clinically affected as well. The affected pups appear stunted and dehydrated, and those surviving the clinical phase have delayed growth and pelage development (Hoover et al., 1985). At necropsy, affected pups consistently have stomachs filled with clotted milk. Typically, the small and large intestines are distended with gas and yellowish fluid, and the other viscera are usually normal in appearance. Microscopic changes are quite minimal. Large numbers of cocci can be seen as aggregates on the surface of villi and villous tips in the small intestine, usually without a significant inflammatory component (Gades et al., 1999). In addition to the intestinal syndrome seen by others, Zhang described retinopathy of prematurity as a descriptor for preretinal neovascularization occurring as a direct consequence of spontaneously occurring neonatal diarrhea (Zhang et al., 1997, 1998). All investigators have emphasized that isolation of an Enterococcus species alone is not diagnostic for neonatal enteropathy. This is so because so many clinically normal rats, perhaps all, once removed from gnotobiotic isolators, carry commensal enterococci, and because biochemically identical isolates may differ markedly in pathogenic potential (Etheridge and Vonderfecht, 1992; Gades et al., 1999). Although virulence factors such as adhesins and cytolysins have received increasing attention, little is known about how such factors are triggered to occur in pathogenic strains, and none have been characterized sufficiently to act as diagnostic criteria (Forbes et al., 1998). Given newer knowledge about the intestinal microbiome, it can be hypothesized that dysbiosis related to antibiotic treatment, or microbial restriction by gnotobiotic derivation, may play a role in the pathogenic potential of this commensal. It is known that once pathogenic strains are isolated in pure culture, Koch’s postulates can be fulfilled by oral or intragastric inoculation of neonatal test rats (Hoover et al., 1985; Etheridge and Vonderfecht, 1992). The possible contribution to this syndrome by rat rotavirus seems not to have been considered in most reports.

D. Pseudomonas aeruginosa (Pseudomoniasis) The association of P. aeruginosa with disease (pseudomoniasis) of laboratory rats and mice has only rarely been encountered as a naturally occurring phenomenon in immunologically competent animals. It may be fairly stated that Pseudomonas is a frequently occurring bacterial contaminant with only weak pathogenic potential for unstressed immunocompetent laboratory rodents. It assumes the status of a significant and serious pathogen in the course of experimental treatments that impair normal host defensive responses. The occurrence of pseudomoniasis as a consequence of such treatments is intensified by the ubiquitous and almost

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universal distribution of the organism in conventional stocks of rodents, other laboratory species, and humans, as well as the ability of the organism to thrive in a variety of inanimate materials outside of mammalian systems that include untreated drinking water, dirty bedding, feces, water bottles, and disinfectant and detergent solutions. In the discussion that follows, information derived from experience with mouse infections has been borrowed, where appropriate for application to the rat, because of the similarity of epizootiology of the infection in these two species and comparatively greater availability of information reported with the mouse. The notoriety of pseudomoniasis in research animals has been primarily established in connection with “early mortality” (3e8 days postirradiation) consequent to lethal whole-body X irradiation in the 750e1000 rad (Gy) range. Experience has indicated that lower dose ranges of X irradiation court infection as well. After irradiation, rats and mice infected with Pseudomonas die earlier and in larger numbers than do uninfected animals (Gundel et al., 1932; Miller et al., 1950, 1951; Gonsherry et al., 1953; Hammond, 1954, 1963; Hammond et al., 1954a,b, 1955; Wensinck et al., 1957; Ainsworth and Forbes, 1961; Flynn, 1963b,c; Woodward, 1963; Hightower et al., 1966; Taffs, 1974). Similarly, supervening pseudomoniasis has complicated burn research in laboratory animals (Ross, 1931; Millican, and Rust, 1960; Millican, 1963; McRipley and Garrison, 1964; Millican et al., 1966; Steinstraesser et al., 2005) and has indeed been used as a model for burn wound contamination in humans (Miyake, 1902; Flynn et al., 1961; Millican, 1963; Teplitz et al., 1964a,b; Lemperle, 1967; Stone et al., 1967; Grogan, 1969; Fox et al., 1970; Nance et al., 1970; Neely et al., 1994; Stevens et al., 1994). Pseudomonas infections in laboratory rodents have been complicated or provoked by other experimental stressors, for example, cortisone injections (Millican et al., 1957), infections with other microorganisms (Flynn et al., 1961; Tsuchimori et al., 1994), antilymphocyte serum (Grogan, 1969), tumors, cold stress (Halkett et al., 1968), neonatal thymectomy (Taffs, 1974), and injection abscesses or surgical procedures (Flynn, 1963b; Wyand and Jonas, 1967; Taffs, 1974; Yamaguchi and Yamada, 1991; Bradfield et al., 1992; Heggers et al., 1992; Dixon and Misfeldt, 1994; Wang et al., 1994). P. aeruginosa infection has been used as a model system for the study of cystic fibrosis (Mahenthiralingam et al., 1994; Johansen et al., 1996; and reviewed by Kukavica-Ibrulj and Levesque, 2008), modulation of resistance by diabetes (Kitahara et al., 1995), mucosal colonization (Pier et al., 1992), and oral vaccines (Cripps et al., 1995). With several exceptions occurring in mice, which include reports of otitis media in closed mouse colonies from which P. aeruginosa was incriminated (Ediger et al.,

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1971; Olson and Ediger, 1972; Kohn and MacKenzie, 1980), no convincing evidence exists to suggest that clinically overt disease syndromes may be induced by Pseudomonas in unstressed immunocompetent laboratory rats and mice, although clinical pseudomoniasis should be recognized as an infectious hazard for immunologically impaired mouse and rat genomic mutants (Johansen et al., 1993; Dietrich et al., 1996). The ecology of P. aeruginosa within animal care facilities is complex and does not depend on animal infection. The organism grows well at ordinary room temperatures and may be introduced by a variety of nonsterile supply fomites, including food, bedding, inadequately sanitized cages, food hoppers, water bottles, bottle stoppers, and sipper tubes (Wensinck et al., 1957; Beck, 1963; Van Der Waay et al., 1963; Hoag et al., 1965). Considerable evidence has accumulated that the organism may be introduced by human contacts (animal care technicians, investigators) even within barrier facilities, primarily by contact with ungloved hands (Wensinck et al., 1957; Van Der Waay et al., 1963). Human carriers appear to be relatively frequent (10% e15% of fecal samples and 90% of sewage cultures) (Hunter and Ensign, 1947; Ringen and Drake, 1952; Lowbury and Fox, 1954; Wilson et al., 1961). The organism has been cultivated from the floor, water faucets, water pipe lines, tap water, cages, water bottles, and sipper tubes within animal rooms (Wensinck et al., 1957; Beck, 1963; Van Der Waay, 1963; Woodward, 1963; Hoag et al., 1965; Trentin et al., 1966). P. aeruginosa thrives in quaternary ammonium disinfectant solutions (Pyrah et al., 1955; Plotkin and Austrian, 1958; Lowbury, 1951) and has been cultured from a number of assumedly sterile parenteral materials (for review, see Flynn, 1963c). It is generally believed that the other floral components of the oropharyngeal and gastrointestinal microbiome of untreated conventional animals are effective in regulating the population dynamics of Pseudomonas in vivo. Repeated animal colony survey data indicate that the ordinary incidence of Pseudomonas carriers is no higher than 5%e20% (Beck, 1963; Hoag et al., 1965; Hightower et al., 1966) in comparison to the near 100% incidence in lethally irradiated rats and mice from the same colonies. Ironically, reduction of these other microbial forms by antibiotic treatments or by establishment of the gnotobiotic state in effect removes their inhibitory effect and facilitates colonization by Pseudomonas to a much higher degree when the animals are exposed to it (Flynn, 1963b; Van Der Waay et al., 1963; Hoag et al., 1965). Pseudomonas appears to localize in the oropharynx, upper respiratory, and rectocolonic tissues in colonized normal animals. It is thought that isolations of P. aeruginosa from deeper reaches of the gastrointestinal tract, that is, duodenum or ileum, represent

organisms being transported through the tract rather than localization or colonization of these sites (Hightower et al., 1966). The organism ascends from the normal areas of localization in vivo to become septicemic after treatments that deprive the host of competence or that induce dysbiosis sufficiently to remove inhibition to invasion. It has been shown that bradykinin and proteases generated in infective foci facilitate septicemic ascension of the organism (Sakata et al., 1996). Septicemia and hematogenous distribution to such organs as the spleen, lungs, and liver generally describe the pathogenesis of pseudomoniasis, and there is some evidence that even peripheral localizations, for example, abscesses of middle and inner ears, pyelonephritis, and skin burn wound sepsis may be hematogenously initiated (Teplitz et al., 1964a,b). The gross pathology therefore is generally that of a septicemia, that is, congestion of splanchnic and thoracic viscera with occasional abscesses of organs with capillary nets. The usual circumstance, however, is that of fulminating infection with onset of death before pyogenic lesion development. Experimentally, a number of factors have been shown to bear on or to modulate the host’s reaction to P. aeruginosa (for review, see Baker, 1998). Heritable differences in susceptibility have been shown with inbred mice, for example, mice of the BALB/c strain are resistant to pulmonary infection but those of the DBA/2 strain are susceptible (Morrissette et al., 1995, 1996). Both humoral and cellular immunity arise and act to modulate the course of experimental infection (Dunkley et al., 1994; Pier et al., 1995; Stevenson et al., 1995) and are enhanced by treatments with vitamin B2 and interleukin (IL) (Araki et al., 1995; Volgels et al., 1995). Cellular components of host resistance include Thelper type 1 cells (Fruh et al., 1995), macrophages and neutrophils (Qin et al., 1995), and tumor necrosis factor (TNF) (Gosselin et al., 1995). Virulence factors associated with P. aeruginosa have been investigated, and it is recognized that Pseudomonas strains vary in virulence (Furuya et al., 1993). Such factors as the bacterial flagellum (Mahenthiralingam and Speert, 1995), pyoverdin (Meyer et al., 1996) and pyocyanin production (Shellito et al., 1992), elastase (Tamura et al., 1992), and exotoxins (Hirakata et al., 1995; Miyazaki et al., 1995; Gupta et al., 1996; O’Callaghan et al., 1996; Pittet et al., 1996; Tank et al., 1996) have been shown to be important elements that determine virulence and have been characterized in the course of model systems of surgical wound infection, respiratory infection, or surgical sepsis. Diagnosis is established by the characteristic case history that usually involves immunosuppressive pretreatment or treatments that induce a shock-like state, and by isolation of P. aeruginosa. It may be isolated from heart

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blood or spleen of septicemic cadavers. The organism is aerobic and grows readily on blood agar, usually producing pigments that darken the agar initially and also a powerful hemolysin that may clear the entire plate. The colonial morphology is irregularly round, 2e3 mm in diameter, usually with a smooth matt surface, butyrous consistency, and floccular internal structure. Colonial variants that include mucoid, rough umbonate, rugose, or smaller coliform-like forms are not uncommon. The organism is a typical Gram-negative rod, 1.5e3 mm in length and 0.5 mm wide. Suspect colonies should be isolated in pure culture and confirmed by tests that characterize the pigment-producing and biochemical properties of P. aeruginosa. Although taxonomy of P. aeruginosa has not been in recent flux, it may be found in the older literature under the names Pseudomonas pyocyanea, Bacterium aeruginosum, Bacterium pyocyaneum, Bacterium aerugineum, Clostridium aeruginosis, and Bacterium pyocyaneus. More typically, however, the organism is suspected in dying animals, or detected by monitoring programs used for microbiologic surveillance of laboratory animals or component equipment. A number of specialized isolation or detection systems for presumptive cultural identification of P. aeruginosa oriented to its biochemical properties have been developed for such purposes. Such studies have emphasized the desirability of enrichment broths for primary isolation followed by subculture to selective or differential semisolid media. Direct inoculation of agar plates for primary isolation from test materials (e.g., water, swabs, feces, or tissue samples)deven when using such recommended media as brilliant green agar, Pseudomonas agar (King’s medium B-Flo agar), glycerol-peptone agar (King’s medium A), or Wensinck’s glycerol agardhave been shown to yield fewer isolations than when the same test materials are selectively enriched in broths before subculture on agar (Hoag et al., 1965). A number of broths for primary culture have been explored, including Wensinck’s glycerol enrichment broth (Wensinck et al., 1957), tetrathionate broth, and Koser’s citrate medium, which may be enriched by addition (10% by volume) of brainheart infusion broth (Beck, 1963; Woodward, 1963). There is some evidence favoring Koser’s citrate medium as the broth of choice for this purpose (Hoag et al., 1965). King’s medium B slants may be used to demonstrate Wood’s light fluorescence of fluorescein pigments diffusing from surface growth of several Pseudomonas species, including P. aeruginosa. There is no convincing evidence to consider Pseudomonas species (other than P. aeruginosa) as pathogens of laboratory rodents, even though they are commonly isolated in the course of health surveillance testing. Presumptive determination of the presence of P. aeruginosa may be made by the observation of water-

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soluble greenish-blue pigments (pyocyanin) in the medium, which may be extracted in chloroform. The latter method, without subculture, has been used for routine screening of water and swabs of water bottles and bottle parts (Flynn and Greco, 1962; Beck, 1963; McPherson, 1963). Other test materials, including oropharyngeal swabs, rectal swabs, feces, and tissue samples, should be subcultured from 24-h broth primary cultures onto agar plates (e.g., brilliant green or glycerol-peptone agars). On brilliant green, P. aeruginosa colonies are reddish and a pronounced red color (nonlactosefermenting, basic pH) develops in the agar. On glycerol-peptone agar, the greenish-blue pigment may be seen diffusing through the medium. Colonies should be confirmed as oxidase positive by the development of a purple color within 15 s when inoculating loop samples are applied to filter paper discs soaked in Kovac’s reagent (Hightower et al., 1966). Final confirmation of individual isolates should be made by establishment of a biochemical profile consistent with this organism (Hansen, 2000). Several systems have been established for the serological (O and H antigens) and pyocin typing of P. aeruginosa (for reviews, see Cowan, 1974; Lennette et al., 1974; Wilson and Miles, 1975; Urano, 1994); however, little correlation between type and pathogenicity or between type and origin of animal isolate has emerged from these studies (Matsumoto et al., 1968; Maejima et al., 1973). These systems and bacteriophage typing are being replaced by pulsed field gel electrophoresis typing (Speert, 2002). Investigations of the method of choice for screening rodent populations have generally established a higher rate of success from the culture of oropharyngeal swabs than fecal samples, and the former is recommended for monitoring programs designed to detect shedder or carrier animals (Trentin et al., 1966). Similarly, screening of water bottle contents after 2e4 days and of water bottle parts have generally yielded higher percentages or indications of carrier animals than do fecal cultures (Beck, 1963; Flynn, 1963a,c; Woodward, 1963; Hoag et al., 1965), although it has been shown that water bottles may frequently fail to yield Pseudomonas from cages with mice having positive oropharyngeal cultures, particularly when the bottles are well sanitized (Trentin et al., 1966). Serological tests have been developed to detect antibodies to P. aeruginosa (Johansen et al., 1993), but their utility for diagnostic surveillance programs has not been explored. It has been established that early detection of pseudomoniasis in human cystic fibrosis was more successful by using a combination of serology (to detect P. aeruginosa antigens) and PCR than microbiologic culture (Filhol et al., 2007). This approach may be of use in animal studies where early invivo detection is critical.

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A variety of approaches has been explored to cope with the problems of pseudomoniasis in immunosuppressed or traumatized animals. Although there are some antibiotics (e.g., gentamicin) (Summerlin and Artz, 1966; McDougall et al., 1967) and bacterocidins (e.g., pyocin) (Merrikin and Terry, 1972) that have been used to extend the life of X-irradiated animals (Wolf et al., 1965), the approach of mass treatments using antibiotics for control of Pseudomonas within rodent colonies has been generally concluded to be ineffective (Hammond, 1963; Hoag et al., 1965; Lusis and Soltys, 1971). Similarly, the use of bacterins for active immunization and passively administered anti-P. aeruginosa hyperimmune sera have been explored and found to have limited value for control of pseudomoniasis in immunosuppressed animals (McDougall et al., 1967; Lusis and Soltys, 1971). The treatment of drinking water with sodium hypochlorite to provide 10e12 ppm chlorine, adjusting water to pH 2.5e2.8 by the addition of concentrated hydrochloric acid, or, preferably, both have been found to suppress Pseudomonas sufficiently to prevent its effects in irradiated or burned animals (Beck, 1963; Woodward, 1963; Hoag et al., 1965). Potable tap water chlorinated to 2 ppm at the source has been found to drop to less than 1 ppm at the tap and to have little inhibitory effect on Pseudomonas (Trentin et al., 1966; Les, 1973). Similarly, chlorine at the 10-ppm level initially is effective in sanitizing water bottles to eliminate Pseudomonas but decreases to 6 ppm by 24 h, 2.5 ppm by 48 h, and 0.25 ppm by 72 h and resembles tap water (McPherson, 1963). Such bottles, with sodium hypochlorite alone, may become recontaminated by 48e72 h if the mice are oropharyngeal carriers of Pseudomonas. It was found earlier that Pseudomonas is quite sensitive to low pH (2.5e2.8) and that chlorinated/acidified drinking water was effective in eliminating the Pseudomonas carrier state (and substantially suppressing coliform and Proteus populations) in laboratory mice (Schaedler and Dubos, 1962). The combination of acidified and chlorinated water is recommended for mass treatment as a standard method for control of pseudomoniasis in immunocompromised animals. Palatability studies have indicated little, if any, preference for untreated tap water compared with chlorineacid-treated drinking water (McPherson, 1963; Thunert and Heine, 1975). However, at least one study has indicated retarded growth on the basis of reduced intake in some mouse strains (Les, 1968a). Several studies through reproductive lifetime cycles have failed to demonstrate deleterious effects of such treatments (Mullink and Rumke, 1971); indeed, most parameters of reproductive performance have increased (Schaedler and Dubos, 1962; McPherson, 1963; Les, 1968b). The low pH has no demonstrable effect on rat tooth enamel

(Tolo and Erichsen, 1969). It is known that a small proportion of rats and mice on treated water may continue to shed Pseudomonas in feces (Hoag et al., 1965; Trentin et al., 1966). If it is desirable to do so, such animals may be detected by fecal or oropharyngeal screening and removed from the colony. The decision to recommend initiation of acidificationechlorination treatment of drinking water in rat colonies resolves itself to an assessment of the risk of pseudomoniasis in the course of experimentation and should be determined by the nature of the research program. No specific disease ameliorative or preventive effects with other defined rat pathogens have been shown for chlorinated/acidified drinking water.

E. Helicobacter Infection The index cases of Helicobacter infections in mice were reported as incidental findings of chronic hepatitis in several inbred strains of mice originating from a Frederick Cancer Center barrier-maintained facility (Fox et al., 1994; Ward et al., 1994a,b). It was determined that the lesions were caused by a newly recognized helical bacterium, later designated Helicobacter hepaticus, isolated from affected livers. The spiral bacterium also colonized the cecal and colonic mucosae of the mice. The agent could be passed to satisfy Koch’s postulates as the incitant of the hepatitis and, as was learned later, of the hepatomas induced as late-stage sequelae in certain mouse strains (Ward et al., 1994a,b). H. hepaticus has not been reported to infect rats. The H. hepaticus cases opened a new chapter in infectious diseases of laboratory animals and served as the introduction to an intensive diagnostic and research effort to characterize the nature and biology of this new genus of gastrointestinal pathogens (for reviews, see Fox and Lee, 1997; Weisbroth et al., 1999a; Whary and Fox, 2004). Based only on morphologic criteria, Helicobacter-like spiral bacteria had been noted earlier in rat ceca (Davis et al., 1972). Later, Phillips and Lee (1983) isolated the spiral-shaped bacteria from intestinal mucosa of rats and mice on a selective medium for campylobacter. Using electron microscopy and biochemical testing, they concluded that the spiral bacterial isolates did not correspond to any known genus (also see references in Solnick and Schauer, 2001). The genus Helicobacter is generally separated into two groups: gastric Helicobacter spp. and enterohepatic Helicobacter spp. according to their preferred site of colonization. Occasionally, enterohepatic groups colonize the biliary tree (Whary and Fox, 2004; Cacioppo et al., 2012a). There are at least seven species of enteropathic Helicobacter reported to naturally colonize rats, Helicobacter bilis, Helicobacter trogontum, Helicobacter muridarum, Helicobacter rodentium, Helicobacter

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TABLE 12.3

Summary of Helicobacter spp. in Rats and Mice.

Helicobacter spp. Helicobacter bilis

471

Speciesa a

R

Infection with Clinical Significance M

Typhlocolitis, proctitis in Cr:NIH-rnu nude rats; no clinical significance in immunocompetent rats Hepatitis, typhlitis in immunocompromised mice

H. ganmani

R

H. hepaticus

M

No clinical significance recognized in rats or mice

M

Hepatitis in multiple mouse strains, hepatocellular carcinoma in A/JCr strain; typhlitis, colitis in scid/NCr

H. muridarum

R

M

Typhlitis and hepatitis in immunocompromised mice; no clinical significance recognized in rats

H. pullorum

R

M

No clinical significance recognized in rats or mice

H. rodentium

R

M

No clinical significance recognized in mice or rats

H. trogontum

R

H. typhlonius

R

M

Typhlocolitis in C.B.-17 scid mice, diarrhea, intestinal bleeding, rectal prolapse in IL-10/ mice

No clinical significance recognized in rats

No clinical significance recognized in rats (PCR identified) R ¼ laboratory rats, M ¼ laboratory mice (Cacioppo et al., 2012a,b; Haines et al., 1998; Johansson et al., 2006; Whary and Fox, 2004). a PCR (þ)/colonized/infected.

typhlonius, Helicobacter ganmani, and Helicobacter pullorum (Table 12.3). However, only H. bilis has been shown to cause clinical disease in rats. Haines et al. (1998) found during a routine necropsy of a retired male nude rat (Cr:NIH-rnu), histological lesions of proliferative typhlitis and the presence of spiral-shaped bacteria within cecal crypts. Subsequent to this initial case, an additional 10 males, 5e8 months old, from the same colony were necropsied. Of the 11 rats, 5 had mild clinical signs consisting of acute perianal inflammation and mild diarrhea. The cecum of each of these 5 rats had focal or diffuse whitish thickened areas. Eight of the 11 had proliferative colitis and proctitis of varying severity. The most severe large intestinal lesions were usually in the cecum. Microscopically, the crypt epithelium was hyperplastic with a decrease in goblet cells. The lamina propria contained a lymphocytic and plasma cell infiltrate, and there was erosion and ulceration of the mucosa with extension of inflammation into the submucosa. Crypts were mildly dilated with bacteria, particularly in the cecum, and Steiner’s staining revealed masses of curved/spiral rod-like bacteria compatible with Helicobacter. No other significant lesions were noted in other major organs, including stomach, small intestine, and liver. Identification of the isolates as H. bilis was achieved by PCR, amplicon sequencing, and negative stain electron microscopy. Using the H. bilis isolate from the foregoing naturally infected rats, the authors were able to reproduce the large bowel disease in H. bilis-free male athymic nude rats. Noteworthy in this report is that only male nude rats developed the inflammatory large bowel disease.

An excellent color illustration of a thickened rat colon may be seen in the text by Barthold et al. (2016). A unique spiral-shaped organism was isolated from the intestinal mucosa of rats and mice by utilization of a campylobacter medium. Experimental infection with this isolate into gnotobiotic rats and mice resulted in its colonization of the intestinal crypts in both species (Phillips and Lee, 1983). This helical bacterium was shown by a genus-specific Helicobacter DNA probe to be a member of that genus, and from subsequent phylogenetic data shown to be a new species named H. muridarum (Lee et al., 1992). Colonization with H. muridarum is not known to be associated with clinical signs or pathology. H. trogontum isolated from the colonic mucosa of Wistar and Holzman rats has not been associated with either overt or clinically inapparent disease in the rat (Mendes et al., 1996), nor has it been reported in natural infection of other rodents. H. pullorum, a species that is a known pathogen in poultry and humans, has been shown to colonize rodents either naturally or experimentally. Surveillance testing of barrier-housed C57BL mice and Brown Norway rats detected a Helicobacter sp. that was later confirmed to be H. pullorum (Cacioppo et al., 2012a). In a study, rats orally dosed with the organism were found to be H. pullorumpositive by fecal PCR for the entire 30week study and to have had an IgG antibody response to the infection. This was reported to be the first study in which an enterohepatic Helicobacter sp. that infects humans also persistently infects rats. Necropsy confirmed that the cecum and colon were the primary

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sites for localization of the organism; however, no intestinal or hepatic pathology was observed in any H. pullorum-colonized rats. Unlike experimental infection in Brown Norway rats, Cacioppo et al. (2012b) found that similarly administered oral doses of H. pullorum to 10 Sprague-Dawley rats resulted in only 2 of 10 rats being persistently infected 12 weeks postinfection. This finding and the inability to successfully transfer H. pullorum by contaminated cage bedding suggests that Sprague-Dawley rats are resistant to infection with the organism and unsuitable for use as sentinels for detection of this Helicobacter. H. ganmani, known to colonize laboratory mice (Robertson et al., 2001), was identified by partial 16S rDNA sequence analysis from cecal and colon samples taken from 40 asymptomatic rats originating from eight different animal facilities in Sweden (Johansson et al., 2006). This was the first report of H. ganmani in naturally occurring colonization of rats. Histopathology remains an important tool for describing morphologic changes attributable to Helicobacter and for confirming clinical diagnosis in lesioned animals, but not as a screening test in clinically normative rodents. Use of silver staining of intestinal specimens is not particularly useful because many other enteric bacterial agents will also be observed with the silver stain (Whary and Fox, 2004). Microbiologic isolation of enterohepatic Helicobacter species is best achieved by filtering homogenized specimens through either a 0.45-or 0.65-mm filter, depending upon the Helicobacter spp., to eliminate enteric commensals. This filtrate is then cultured on blood agar plates supplemented with trimethoprim, vancomycin, and polymyxin to inhibit contaminating organisms from clinical materials (tissue or fecal suspensions). Plates are incubated at either 37 or 42
C in a specialized microaerophilic incubation environment (nitrogen, hydrogen, CO2 in a ratio of 90:5:5) for 3e7 days (Shames et al., 1995; Whary and Fox, 2004). However, plates should be kept for 21 days to ensure slowly replicating Helicobacter colonies are observed before discarding. Greater bacterial numbers can be achieved by inoculation of plate isolates into Brucella broth containing 5% fetal bovine serum and incubation for 24e48 h on a rotary shaker. Isolated Helicobacter cultures are identified by their spreading growth films with spiroform Gram-negative bacteria. As is true for all bacteria, isolation in pure culture is required to enable speciation of isolates on the basis of biochemical criteria (see tables in Mendes et al., 1996; Fox and Lee, 1997; Whary and Fox, 2004). Of the Helicobacter spp. that colonize rats, H. bilis, H. muridarum, and H. trogontum are catalase and urease positive; H. pullorum, H.rodentium, and H. ganmani are catalase positive and urease negative. When strains are isolated that do not match existing profiles, a new species may have

been encountered (Dewhirst et al., 2000b). In summary, culture has been widely used as a screening method, but important limitations include the obscuring presence of contaminating microbial overgrowth and low Helicobacter populations, both of which bear on the reliability of negative results (Weisbroth, 1999b) (Table 12.4). Most of what is known about the immunological response to Helicobacter infection is based on investigative experience with mice. The function the immune response has in either limiting colonization or in contributing to the intensity of the inflammatory response to Helicobacter infections that can progress to cancer in mouse models has been widely investigated (Whary and Fox, 2004). The immune response to Helicobacter is prompt and appears proportional to the intensity of infection or degree of tissue invasiveness but may not confer protection (Fox and Lee, 1997). Unlesioned and naturally resistant mouse strains may be colonized by H. hepaticus without development of a detectable antibody response (Ward et al., 1996). After experimental infection of Brown Norway rats with H. pullorum, IgG seroconversion measured by ELISA occurred in four of six colonized rats at 2weeks postinfection. A fifth rat seroconverted at 12 weeks, whereas the single uncolonized rat did not seroconvert (Cacioppo et al., 2012b). Both whole-cell sonicates (Fox and Lee, 1997) and outer cell membrane proteins (Livingston et al., 1997; Whary et al., 1998) have been used to prepare ELISA antigens. In parallel with lesion development and serum alanine transaminase levels, antibodies have first been detected in naturally infected animals (of susceptible strains) from enzootically infected colonies at about 6 months of age. The intensity of the optical density of the ELISA titers increases with age, reaching maximal levels at about 12e18 months (Fox and Lee, 1997). The ELISA test using H. hepaticus antigens appeared quite specific, recognizing antibodies to H. hepaticus but not to H. bilis or H. muridarum (Livingston et al., 1997). Serology has received some attention as a screening method (Whary et al., 2000a) but, similar to cultural isolation, has important limitations that inhibit its endorsement as a generally useful diagnostic tool. Because antibody levels to Helicobacter are generally proportional to the intensity of microbial challenge, positive serology is probably reliable as an indicator of exposure to antigens used in the test. The problem is that negative serology is not a reliable result for any of a number of reasons, including (1) innate host resistance that prevents lesion development or significant antigenic challenge; (2) initially low populations of Helicobacter in the gastrointestinal reservoir; (3) serum samples that may be drawn early in the course before development of detectable antibody levels; and, perhaps most importantly, (4) the possibility of false negative because of infection by Helicobacter species that stimulate antibodies not recognizing antigens used in the test (i.e.,

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TABLE 12.4

Diagnostically Important Pasteurellaceae in Laboratory Rats and Mice. Pathogenic Rats a

Mice b

ID

IC

ID

IC

Surveillance/Diagnostic Schedulec

Pasteurella pneumotropicus

þ/

þ/e

þ

þ/

Monitor every 3 months

Pasteurella heylii

þ/

þ/e

þ

þ/

Monitor every 3 months

Muribacter muris

e

e

þ/

þ/

Dependent on research

Haemophilus spp. (V-factor dependent)

þ/e

þ/e

Dependent on research

Pasteurellacae (V-factor dependent)

e

e

Dependent on research

þ/ ¼ Pasteurellaeae are characteristically nonpathogenic to weakly pathogenic in most cases, except in immunodeficient mice. However, in the presence of copathogens such as Mycoplasma pulmonis, CAR bacillus, or Sendai virus, they may contribute to the observed pathology in rats and mice. a Immunodeficient/impairment. b Immunocompetent. c FELASA recommendations. Adhikary, S, Bisgaard, M., Dagnaes-Hansen, F. and Christensen, H. (2017a). Clonal outbreaks of Pasteurella pneumotropica biovar Heyl in two mouse colonies. Lab. Anim. 51, 613e621; Adhikary, S., Nicklas, W., Bisgaard, M., Boot, R., Kuhnert, P., Waberschek, T., Aalbaek, B.Korczak, B., and Christensen, H. (2017b). Rodentibacter gen. nov. including Rodentibacter pneumotropicus comb. nov., Rodentibacter helylii sp. nov., Rodentibacter myodis sp. nov., Rodentibacter ratti sp. nov., Rodentibacter heidelbergensis sp. nov., Rodentibacter trehalosifermentans sp. nov., Rodentibacter rarus sp. nov., Rodentibacter mrazii and two genomospecies. Int. J. Systemat. Evolutionary Microbiol. 67, 1793e1806. (First published online: 20 June 2017); Boot, R., van den Berg, L., Van Lith, H.A. and Veenema, J.L. (2005). Rat strains differ in antibody response to natural Haemophilus species infection. Lab. Anim. 39, 413e420; Boot, R., Nicklas, W., Christensen, H., 2018. Revised taxonomy and nomenclature of rodent Pasteurellaceae: implications for monitoring. Lab. Anim. 52, 300e303; Hayashimoto, N., Yasuda, M., Ueno, M., Goto, K., and Takakura, A. (2008). Experimental infection studies of Pasteurella pneumotropica and V-factor dependent Pasteurellaceae for F344-rnu rats. Exp. Anim. 57, 57e63; Mahler, M., Berard, M., Feinstein, R., Gallagher, A., Illgen-Wilcke, B., Pritchett-Corning, K., Paspa, M., 2014. FELASA recommendations for the health monitoring of mouse, rat, hamster, Guinea pig and rabbit colonies in breeding and experimental units. Lab. Anim. 48(3), 178e192; Miller, M., Zorn, J., and Brielmeier, M. (2015). High-resolution melting curve analysis for identification of Pasteurellaceae species in experimental animal facilities. PLoS One. 10(11). doi.10.1371/journal.pone.0142560; Nicklas, W. (1989). Haemophilus infection in a colony of laboratory rats. J. Clin. Microbiol. 27, 1636e1639; Towne, J.W., Wagner, A.M, Griffin, K.J., Suntzman, A.S., Frelinger, J.A. and Besselsen, D.G. (2014). Elimination of Pasteurella pnemotropica from a mouse barrier facility by using a modified enrofloxacin treatment regimen. J. Amer Assoc Lab Anim Sci. 53(5):517e522).

issues of specificity). One practical problem is that validated antigens are not generally available for utilization in testing programs. Diagnosis of Helicobacter colonization in the rat is best accomplished by using genus- and species-specific PCR assays. The PCR is the most useful single screening test for detection of murine helicobacters from clinical materials, including tissue and fecal specimens (Battles et al., 1995; Shames et al., 1995; Beckwith et al., 1997; Riley et al., 1997; Hodzic et al., 2001) and retrospective material, including wet, fixed tissues, paraffin-embedded tissues, and even stained histologic specimens. The method is based on the detection of unique and subsequently amplified 16S rRNA gene sequences of the agent extracted from the sample material. The test is rapid (1 day) and, by use of restriction endonucleases, can be used to speciate Helicobacter DNA detected in generic tests (Riley et al., 1996). The procedure is specific toone or more Helicobacter sequences specified by the primers used for amplification and is not complicated by the presence of contaminating microorganisms in the sample material. One important property of the PCR test is extreme sensitivity for detection of low numbers of Helicobacter in the sample specimens. In direct comparison, PCR tests detected as positive 30%e50% more specimens found negative from animals also tested by culture

and/or histopathology or electron microscopy (Shames et al., 1995). Furthermore, the method permits sample pooling up to dilutions of 1:5e1:10 of positive material without significant loss of sensitivity, thus economizing on the cost of testing (Whary et al., 2000a,b). The fluorogenic nuclease PCR assay has been shown to have several advantages over gel-detection PCR for detection of Helicobacter spp. H. hepaticus, H. bilis, and H. typhlonius (Drazenovich et al., 2002). This assay method eliminates the need for post-PCR processing, may provide a greater specificity, and is able to generate quantitation of bacterial load. The combination of PCR and serology for screening programs appears to offer superior detection rates (Whary et al., 2000a). Because of concern about the potential of Helicobacter infections to spread within both breeding and user facilities and to adversely impact research programs, treatment modalities have been explored as a means of eradicating Helicobacter from asymptomatic-infected carriers and of reducing the clinical manifestation in susceptible mouse strains. It has been shown that 2-week treatments of amoxicillin alone, dosed via the drinking water, were effective in eliminating (or preventing) H. hepaticus infection in weanlings but not in older mice with established enteric colonization (Russell et al., 1995). Triple treatment with combinations of

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amoxicillin, metronidazole, and bismuth (known to eradicate Helicobacter pylori in humans and Helicobacter mustelae in ferrets) given by gavage three times daily for 2 weeks appeared effective in eradicating H. hepaticus infections in 6- to 8-month-old susceptible inbred mice (Foltz et al., 1995), but the labor-intensive nature of the dosing regimen limits usefulness of this individual treatment. More promising was the ad libitum oral treatment with amoxicillin triple treatment formulated into flavored dietary wafers used to eradicate H. hepaticus in 6- to 10-month-old naturally infected susceptible inbred mice (Foltz et al., 1996). As with the gavage method, the dietary wafer modality appears most useful to clear small, defined groups of the infection. Both modalities share the general problem with antibiotic treatments to suppress rather than to fully eradicate, thus setting the stage for eventual rebound of residual Helicobacter populations after withdrawal of treatment. An additional area of concern is the uncertain susceptibility of other Helicobacter species to the treatments discussed earlier for H. hepaticus. It has been reported (Shomer et al., 1998) that although diarrheas caused by dual infection by H. bilis and H. rodentium (in scid mice) could be clinically ameliorated, the Helicobacter infections were not eradicated after 2-week treatments with dietary wafers. There is limited information on the efficacy of antibiotic modalities for eradication of Helicobacter spp. in rats. In one report (Jury et al., 2005), a medicated feed formulation containing 0.07 mg omeprazole, 3.3 mg metronidazole, 6.7 mg amoxicillin, and 1.7 mg clarithromycin, incorporated into a 5-g nutritional complete grain-based, baconflavored tablet (Bio-Serv, Frenchtown, NJ), was used in two feeding protocols. In the first, PCR-positive, pregnant, inbred Brown Norway rats were started on day 7 of gestation and kept on the feed until the pups were weaned at 3 weeks of age. Fecal samples were taken at 1, 3, and 6 months postweaning, all samples tested Helicobacter spp. negative at all time points. In the second feeding protocol, another inbred rat strain, Flinders Sensitive Line (FSL) adult male rats, were fed the medicated feed for 2 weeks, fed nonmedicated feed for 2 weeks, and given the medicated feed again for a 2-week cycle. The FSL rats were known to be infected with H. bilis and H. rodentium. These rats tested negative for Helicobacter spp. after two medication cycles and remained free through the end of the 8-month study. Not a great deal is known about other means to eradicate Helicobacter infections. Successful eradication of Helicobacter infections should be oriented to principles familiar with eradication of other highly infectious murine pathogens, such as depopulation (or rederivation) of affected units, sanitization of empty rooms and

repopulation with the same strain stocks rederived by cesarean section or embryo transfer, or use of stocks free of Helicobacter brought in from the outside. Stringent maintenance regimens for admission of personnel and reprocessed equipment and supplies should be put in place to prevent reinfection of the clean stock. A number of qualifiers bear on the pathogenic significance of rodent Helicobacter infections (Chichlowski and Hale, 2009). It would seem warranted to regard the rodent helicobacters as essentially opportunistic enteric bacteria with only low-grade potential to cause disease. That said, reports are steadily accumulating that detail enterohepatic disease syndromes in variably immunocompetent but susceptible inbred mice, as well as in immunodeficient mouse and rat strains. Complicating the issue is the proliferating diversity of microbial species and strains within the genus Helicobacter. In general, the Helicobacter species are fairly host specific, but not entirely. For example, both H. bilis and H. muridarum of the mouse have been isolated from rats (Fox and Lee, 1997), and H. bilis has been reported as a cause of enteric disease in athymic rats (Haines et al., 1998). Most laboratories working with these agents find occasional isolates with profiles that do not neatly match one of the characterized species, thus not all of the variants have been carefully speciated as yet (see Dewhirst et al., 2000b). The Helicobacter species also vary in pathogenicity, even in susceptible hosts. ThusH. bilis and H. hepaticus have been documented as pathogens on a number of occasions in a wide number of mouse strains, as well as H. bilis as a pathogen of immunodeficient rats, and should therefore be regarded as frank pathogens of susceptible hosts. On at least two occasions (Phillips and Lee, 1983; Lee et al., 1992), H. muridarum (a species noted earlier as isolated from rats) has been microscopically associated with mild gastritis in naturally infected mice. There is no strong, convincing evidence, at present, to indict H. trogontum as a pathogen of laboratory rats under circumstances of natural infection. Nonetheless, the (tenuous) designation of “nonpathogen” should be cautiously applied. Given the propensity of this genus to cause enterohepatic disease in such a widely diverse range of mammalian hosts, it may well only be that the right mix of agent/host/milieu has not been encountered to eventuate and be recognized as a cause of clinical disease. A good example of this potential was reported in which scid mice developed hypertrophic colitis with diarrhea when dually infected with H. bilis and H. rodentium. When infected with H. rodentium alone (a species previously not known to be pathogenic), typhlocolitis was caused experimentally in the same strain host mice (Shomer et al., 1998).

CLINICAL CARE AND DISEASE

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F. Clostridium piliforme Infection (Tyzzer’s Disease) Tyzzer’s disease of the rat, caused by Bacillus piliformis, occurs as a latent and asymptomatic condition, or may be an acute and generally fatal epizootic with cardinal clinicopathologic features of megaloileitis, focal necrosis of the liver, myocarditis, and diarrhea. The condition is named after Ernest Tyzzer of Harvard, who initially reported the disease as a fatal epizootic of Japanese waltzing mice, Mus bactrianus (Tyzzer, 1917; Kerrin, 1928). Tyzzer’s original report of the disease, with descriptions of its histopathology, still stands as the fundamental study. The disease has emerged, with worldwide distribution, as one of the major infectious diseases of the rat. Naturally occurring disease has been described in the laboratory rat (Fujiwara et al., 1963a; Takagaki et al., 1968; Stedham and Bucci, 1969, 1970; Jonas et al., 1970; Yamada et al., 1970). It has been observed preeminently in the laboratory mouse (Gard, 1944; Rights et al., 1947; Porter, 1952; Saunders, 1958; Carda et al., 1959; Tuffery, 1956; Friedmann et al., 1969; Kaneko et al., 1969; Peace and Soave, 1969; Francis, 1970; Meshorer, 1974), and also in a number of other mammalian species, including the gerbil (Carter et al., 1969; White and Waldron, 1969; Veazey et al., 1992), hamster (Fujiwara et al., 1968; Takagaki et al., 1974; White and Waldron, 1969), guinea pig (Boot and Walvoort, 1994), rabbit (Allen et al., 1965; Cutlip et al., 1971; Ganaway et al., 1971b; Van Kruiningen and Blodgett, 1971), and larger animals such as primates (Niven, 1968), cats (Kovatch and Zebaith, 1973; Ikegami et al., 1999a), red pandas, calves (Ikegami et al., 1999b), horses (Hall and Van Kruiningen, 1974; Thomson et al., 1977; Hook et al., 1995; Fosgate et al., 2002), and immunocompromised humans (Smith et al., 1996). Ganaway et al. (1971a) pointed out that only two articles had been reported on the disease between 1917 and 1933 and six more articles between 1933 and 1959. Of 35 articles between 1959 and 1971, 23 had appeared since 1965. In that same context, the disease was recognized only in Mus musculus, M. bactrianus, and their crosses before 1965, whereas naturally occurring Tyzzer’s disease has since been recognized in a wide range of other animals. It is uncertain if the disease is emerging or, rather, being subjected to more intensive surveillance. Certainly, the advent of generally available serologic reagents appeared to demonstrate the common occurrence of serologic positivity in rodent health surveillance programs, especially the seeming occurrence of C. piliforme infection in the absence of Tyzzer’s disease (Hansen et al., 1990, 1994; Motzel et al., 1991; Hansen and Skovgaard-Jensen, 1995) although the reliability of serologic detection as indicative of actual C. piliforme infection is widely refuted (see ahead). The degree to which this

475

disease has interfered with the interpretation of experimental results is difficult to assess for several reasons; however, it is thought to be substantial. On the one hand, it is likely that the disease frequently has gone unrecognized by investigators, and on the other hand it is a general fact that ruined experiments are seldom published (for documented experimental complications, see references in Lindsey et al., 1991). In England, Tyzzer’s disease was claimed to have ruined more carcinogenesis experiments than has any other single disease (Wilson and Miles, 1975). The taxonomic status of C. piliforme, the Tyzzer’s disease agent, has been greatly clarified since the first edition of this text. Wilson and Miles (1975) proposed the name Actinobacillus piliformis based on the morphologic analogy of monilial thickenings at irregular locations in the vegetative rods seen in Streptobacillus (Actinobacillus) moniliformis (muris). The analogy appears to end there, however, and the name originally proposed by Tyzzer (B. piliformis) had been retained until recently. On the basis of 16S rRNA sequence analysis, the agent has been assigned to the clostridia and given the name C. piliforme (Duncan et al., 1993). The organism stains poorly with hematoxylin, although it is generally recognizable, with some experience, in hematoxylin and eosin-stained tissue sections. Methylene blue or Giemsa stains are satisfactory for demonstration of the organism in smears and tissue sections. The silver impregnation stains (e.g., Warthine Starry or GMS) similarly are satisfactory and may be the stains of choice for demonstration of the organism in tissue sections. The organism is periodic acideSchiff (PAS) positive and Gram (Brown and Brenn) negative. Excellent color illustrations may be found in the publications by Jonas et al. (1970), Fujiwara et al. (1963a), and Barthold et al.(2016). The morphology of the organism is pleomorphic, but general agreement has been reached that the vegetative phase of long slender rods, approximately 0.5  8e10 mm (or more), are the obligately intracellular forms seen in large numbers (sheaves or bundles) within infected cells. Spore-likemonilial thickenings are seen less frequently in tissue sections from liver and intestine, but more frequently in yolk sac preparations from infected embryonating chicken eggs (Craigie, 1966a). Although Tyzzer believed the organism to be nonmotile, observation of motility by phase-contrast microscopy (Allen et al., 1965; Craigie, 1966a), supported by demonstration of peritrichous flagella using electron microscopy (Fujiwara et al., 1968; Jonas et al., 1970; Kurashina and Fujiwara, 1972), has led to its being accepted as motile. It is generally concluded that C. piliforme may not be successfully cultivated in cell-free media (Thunert, 1984). The two reports to the contrary (Kanazawa and

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Imai, 1959; Simon, 1977) have not been supported by other investigators. The organism may be propagated in embryonating eggs by yolk sac inoculations of 10- to 12-day chick embryos (Allen et al., 1965; Craigie, 1966a; Fries, 1977b) without loss of pathogenicity. Preferentially, the organism may be cultivated in a number of primary and continuous cell lines, for example, mouse fibroblast 3T3, mouse L-929, mouse hepatocyte NCTC 1469, and human carcinoma Caco-2 (Franklin et al., 1993; Kawamura et al., 1983a,b; Riley et al., 1990; Spencer et al., 1990). The viability of the vegetative phase, that is, the bundles of long (8e40 mm) slender rods seen in infected cells, is extremely unstable (Fujiwara, 1967). Infective preparations made from heavily infected livers, even when injected into cortisone-prepared substrate hosts, are likely to fail if made from a donor animal that has been dead more than 2 h. Preparations made from infected yolk sacs lose substantial activity after 15e20 min at room temperature and even more at 37
C. The vegetative phase is killed by heating to 45
C for 15 min. Infectivity is completely lost after 24 h at 4
C. Preparations of the vegetative form may be stabilized by freezing at 70
C. The spore form is considerably more stable. Spores are about 0.5 wide  3 mm long and occur as terminal endospores of vegetative cells or may be free (Riley et al., 1994). They will survive 56
C temperatures for 1 h and repeated cycles of freeze and thaw, but they will not survive 80
C or greater for 30 min or more (Ganaway, 1980). Spores are resistant to ethanol, phenolic germicidal detergents, aldehydes, and quaternary ammonium disinfectants, but spores are inactivated by iodophores, formalin, peracetic acid, and sodium hypochlorite solutions (Ganaway, 1980; Itoh et al., 1987a,b; Hansen et al., 1990, 1992b; Riley et al., 1994). Infectivity of spore forms may be retained in dead yolk sacinoculated chicken eggs for 1 year at room temperature (Craigie, 1966a) and for similar periods in infected mouse and rabbit bedding at room temperature (Tyzzer, 1917; Allen et al., 1965; Christensen, 1968). Natural infections are initiated by oral ingestion of spore forms from the bedding contaminated by infected animals shedding spores in the feces. Both Tyzzer (1917) and Gard (1944) used contaminated cages for transmission studies. Transmission studies directly exploring the oral route have produced varying results. Oral infections in the young rat (Jonas et al., 1970; Kurashina and Fujiwara, 1972), rabbit (Allen et al., 1965), and gerbil (Carter et al., 1969) have generally been successful, with characteristic production of both intestinal and liver lesions. In the mouse, Gorer (1947) used dried liver blocks as a means of stabilizing the sporulated form, which were ground up and placed in cooked cereal for oral transmission studies. Interestingly, Fujiwara et al.

(1973a) reported higher rates of success with raw liver blocks than with ground-up liver suspensions and proposed that cannibalism might serve for transmission and maintenance of the cycle in breeder colonies. Tyzzer’s disease was induced via gastric intubation of untreated suckling mice up to 6e7 days old, with enhanced transmission in cortisone-treated weanlings and adults starved 24e48 h before infection (Fujiwara et al., 1973a,b). It has been suggested that variability in transmission studies may relate to fragility of the vegetative form in vitro and/or failure to administer sufficient spores for oral transmission (Ganaway et al., 1971b). Support for this view was shown in the higher infectivity rate obtained in nonfasted adult mice inoculated with 107 organisms directly into the duodenum or cecum (38% and 50%, respectively) than intragastrically (13%), and also by the demonstration that the infectivity rate was dose related (Fujiwara et al., 1973a,b; Takagaki et al., 1974). Experimentally, it has been demonstrated that spores are shed for 1e2 weeks after oral infection and that rodents exposed to contaminated feces can contract Tyzzer’s disease (Waggie et al., 1984; Itoh et al., 1988; Franklin et al., 1994). Little is known about antecedent factors predisposing the pathogenesis and history of natural infections. Clinically apparent Tyzzer’s disease may appear in epizootic proportions with seemingly little in the way of predisposing stress, but more commonly the disease appears to be precipitated into overt, but sporadic, clinically apparent disease by a number of factors that impair immunocompetence of carrier hosts. Precedent experimental treatments that include tumor transplantation (Tyzzer, 1917; Craigie, 1966b; Takenaka and Fujiwara, 1975), administration of steroid or other immunosuppressive drugs (Takagaki et al., 1963; Fujiwara et al., 1964b; Niven, 1968; Yamada et al., 1970; Taffs, 1974; Fries, 1979a), leukocyte injections (Taffs, 1974), and X irradiation (Takagaki et al., 1963, 1966; Taffs, 1974) have been either observed in the course of natural outbreaks or used to facilitate transmission studies with the Tyzzer’s agent. Other factors bearing on susceptibility are known (or believed) to include poor sanitation and overcrowding (Gard, 1944; Rights et al., 1947; Peace and Soave, 1969), transportation stress (Peace and Soave, 1969), nutritional status (Maejima et al., 1965), dietary form (Gard, 1944), and carbon tetrachloride liver injury (Takenaka and Fujiwara, 1975). In assessing the issue of genetic susceptibility and resistance, Gowen and Schott (Gowen and Schott, 1933; Takagaki et al., 1966) were unable to relate resistance in M. musculus, M. bactrianus, and their crosses to the waltzing factor, dominant white, or sex; they concluded that resistance was best interpreted as depending on a major dominant factor. It has been reported that mouse strains with impaired

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immunocompetence have heightened susceptibility and more severe lesions than do their normative counterparts (Waggie et al., 1981; Livingston et al., 1996). Resistant mouse strains (C57Bl) depleted of natural killer cells, neutrophils, and macrophages have increased susceptibility that is comparable to susceptible strains (DBA/2) (Van Andel et al., 1997). Mice experimentally infected with either toxigenic or nontoxigenic C. piliforme isolates respond within a day with elevation of hepatic TNF-a, interferon (IFN)-g, and serum proteins (Van Andel et al., 2000a) and with elevation of IL-6 (Van Andel et al., 2000b) and IL-12 (Van Andel et al., 1998). Immunologic neutralization of reactive IL-12, in vivo, was associated with increased severity of experimental infections. The latter result was interpreted as indicating a protective role for IL-12 as a mediator of infection. In the rat, it has been shown epidemiologically that certain haplotypes were associated with resistance to natural infection, whereas others were more susceptible (Hansen et al., 1990, 1992a). The balance of evidence now suggests that the pathogenesis of natural infections is in accord with the following sequence: oral ingestion of infectious spores from contaminated litter with establishment of primary infection in tissues of the jejunum, ileum, and cecum. The initial stage is followed by ascension of organisms via the portal vein to the liver and by bacteremic embolization to other tissues, for example, the liver and myocardium. Natural and experimental oral infections in the rabbit, rat, gerbil, and hamster characteristically include intestinal localization of the organism with lesion production in the intestine, as well as liver and (more variably) myocardial lesions (White and Waldron, 1969; Jonas et al., 1970; Yamada et al., 1970; Ganaway et al., 1971b; Van Kruiningen and Blodgett, 1971; Nakayama et al., 1975; Lee et al., 1976; Waggie et al., 1984; Yokomori et al., 1989; Veazey et al., 1992). In natural and experimental oral infections in the mouse, intestinal lesions are more variable and liver lesions do not appear to depend on prior development of intestinal lesions (Fujiwara et al., 1973a,b). The naturally heightened susceptibility of gerbils may account for more extensive lesions in that species, including encephalitis (Veazey et al., 1992), although factors (e.g., variable susceptibility of gerbil stocks and/or pathogenicity of C. piliforme strains) have been suggested as explanations for why gerbils do not always contract the infection (Motzel and Riley, 1992). Virulence determinants have been studied as a function of diversity between C. piliforme isolates. Experimentally, it has been shown (in Caco-2 cultures) that isolates vary in their ability to exit from intracellular entry phagosomes to replicate in the cytoplasm (Franklin et al., 1993). Similarly, isolates vary in their capacity to produce cytotoxins, and toxigenic isolates have been

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shown to exert increased pathogenicity in infected hosts (Riley et al., 1992). Thus the severity of outbreaks may depend on virulence of the isolate, as well as on innate resistance of the host. Bacteremia occurs routinely subsequent to induction of liver lesions (Takagaki and Fujiwara, 1968) and elevation of serum transaminases (ALAT, ASAT) indicative of liver destruction (Hoag et al., 1965; Naiki et al., 1965). Experimental infections of any host species given by parenteral routes (intravenous, intraperitoneal, intracerebral) typically induce lesions of the liver, myocardium, or brain, but not intestinal lesions (Rights et al., 1947; Fujiwara et al., 1964a; Onodera and Fujiwara, 1970, 1972). Few clinical signs specific to naturally occurring Tyzzer’s disease in the mouse and rat have been reported. Even in the presence of enteritic lesions, signs of diarrhea have been variable and not often observed in the rat, although diarrhea is more frequent in other species. Affected rats are depressed, with ruffled haircoat and short period of illness. Infected adolescents may have a potbellied appearance. The few outbreaks that have been studied (before development of high-quality serologic reagents) have indicated low morbidity but high mortality in rat groups. It has been mentioned previously that the serum transaminases are elevated in sick animals. In the mouse, both morbidity and mortality may be high; Tyzzer lost his entire colony of 79 animals (Tyzzer, 1917). Gross lesions in the rat are suggestive and include flaccid segmental dilatation of affected portions of the intestines (3e4 times normal diameter) with an edematous, atonic ileum (Jonas et al., 1970). This gross lesion (megaloileitis), with its in vivo potbellied appearance, may be seen as one of the earliest signs of an outbreak in newly contaminated rat breeding colonies (Hansen et al., 1992a,1994; Hansen, 1996). Ileal lesions may extend to adjacent cecum and jejunum. On the basis of these lesions, Jonas et al. (1970) suggested that several earlier reports of megaloileitis or segmental ileitis in the rat (Geil et al., 1961; Hottendorf et al., 1969) may have represented unrecognized instances of Tyzzer’s disease (although it has been pointed out that the lesion is not pathognomonic and may be induced by chloral hydrate alone (Fleischman et al., 1977). The liver may have many disseminated pale foci, up to several millimeters in diameter, scattered throughout the parenchyma (Jonas et al., 1970; Yamada et al., 1970), but frequently in the rat, it is not lesioned. Circumscribed, grayish foci may be seen in the myocardium in some cases. The mesenteric lymph nodes are usually swollen. No other lesions are seen at necropsy. Excellent color illustrations of gross lesions may be seen in the report by Jonas et al. (1970), in the presentation by Clifford (2012), and in the text by Barthold et al. (2016). Definitive diagnosis of Tyzzer’s disease in the rat continues to rest on histopathologic demonstration of the

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organism in infected tissues; although it has been pointed out that C. piliforme may be isolated from infected tissues and propagated via yolk sac inoculations in chick embryo or permissive cell lines, and these isolates may be used to satisfy Koch’s postulates (Allen et al., 1965; Craigie, 1966a; Franklin et al., 1994). Intestinal lesions are characterized microscopically by transmural involvement of the intestinal wall, particularly the ileum, in a process varying from segmental sharply demarcated necrosis to acute and chronic inflammatory infiltration of ball levels that include the mucosa, submucosa, and muscular and serosal layers. Edema and hemorrhage are common in the subserosal and muscular layers. C. piliforme may be found, often profusely, within epithelial cells of the villi and their crypts. The organism has not been seen deep in the mucosa (Jonas et al., 1970). Although Jonas et al. (1970) described villi as intact but blunted, Japanese investigators have (without detailed microscopic descriptions) described the intestinal process in the rat as an ulcerative enterocolitis (Yamada et al., 1970). Myocardial lesions, more common in the rat than hepatic, vary in size from only several myofibers to complete transmural involvement. Inflammatory infiltrates are variable, and the myocardial process has been described as degenerative rather than necrotic. C. piliforme may be demonstrated by special stains within affected myocardial muscle cells. Microscopic lesions in the liver are initiated by foci of coagulative necrosis varying in size from several micrometers to several millimeters, with a tendency to centrilobular orientation in association with central veins and distinct demarcation between necrotic and adjacent normal hepatic parenchyma. C. piliforme typically is found in viable hepatocytes on the periphery of necrotic foci. Necrotic foci thought to be early developing are relatively acellular and barely eosinophilic, with hepatic cell ghosts and karyolitic debris (Jonas et al., 1970). Lesions interpreted to be later in development demonstrate moderate to marked infiltration with neutrophils, Kupffer cells, macrophages, and fibroblasts, which may almost obliterate previously acellular foci of necrosis (Ganaway et al., 1971a). Mineralized material similar to that seen in rabbit lesions has been seen in necrotic foci of rat livers (Jonas et al., 1970). Giant cells and histiocytes may aggregate peripheral to infiltrated necrotic foci (Jonas et al., 1970). In any given liver, relatively acellular “early” foci may be seen, along with infiltrated “later” foci; others may be seen with granulomatous fibroplasia and giant cells. These observations are suggestive of successive showers of organisms from the intestines. The ultrastructure of hepatic lesions has been described (Fujiwara et al., 1963a, 1968; Jonas et al., 1970; Kurashina and Fujiwara, 1972).

When discussing diagnosis, it needs to be recognized that one of the currently most difficult and confounding diagnostic issues with Tyzzer’s disease of the rat is the frequent occurrence of positivity in colonies monitored by serologic health surveillance programs (see prevalence in Schoondermark-van de Ven et al., 2003) in which it has not been possible to confirm serodiagnosis by a combination of immunosuppressive provocation, traditional histopathology, and molecular techniques. The phenomenon has been widely recognized (Hansen, 1996; Riley et al., 1994; Weisbroth, 1999b). Even FELASA has suggested caution by stating that positive serologic reactions occur frequently in the absence of clinical signs (Nicklas et al., 2002). Initially, the wide use of serologic screening techniques in health surveillance programs (Fries, 1978, 1979b; Motzel and Riley, 1991; Motzel et al., 1991; Hansen et al., 1992b, 1994; Boivin et al., 1993; Hansen, 1996) appeared to present an economical and convenient means for detection of C. piliforme in representative serum samples from rodent populations. However, the common occurrence of serologic positivity in the absence of confirmatory evidence (history, clinical disease, PCR detection of microbiologic genomic material) has led to critical evaluation of serologic methodology. In particular, it has been both demonstrated and generally concluded that antigenic reagents that utilize complex preparations of organisms’ whole cells or their extracts in fact cross-react with antibodies to commensal bacteria carried by test rodents and lagomorphs (Feldmann et al., 2006; Pritt et al., 2010). In this context, the reliability of serologic negatives does probably indicate the absence of C. piliforme antibodies in the test animals, but serologic positives are sufficiently likely to be false positive and require confirmation by other means (gross and microscopic lesions, PCR) to be accepted as diagnostic of infection. It is also true that the era of serologic detection of widespread positivity also coincided with the seeming disappearance of clinical Tyzzer’s disease. There have been no reports in the literature of actual clinically overt infection (in distinction to experimental studies) in laboratory rats in the United States since 1974 (Taffs, 1974), or in Europe since 1994 (Hansen et al., 1992a, 1994) or in mice since 1987. Surely this is a result of widespread availability of gnotobiotically derived rodents. The introduction of molecular methods (PCR) for genomic detection of the organism (Duncan et al., 1993; Goto and Itoh, 1996; Furukawa et al., 2002) has greatly enlarged the dimensions of infection and enabled the distinction to be made between asymptomatic C. piliforme infection and overt Tyzzer’s disease with clinical signs and lesions. Some caution in the interpretation of results is required here as well. Unqualified PCR results may be subject to false positive, depending on the specificity of primers used for the purpose (Feldman

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et al., 2006). Certainly, experimental infections of immunocompetent rat and mouse hosts support the concept of an acute, transient, immunogenic, but self-limiting infection that may last no more than 1e2 weeks (Waggie et al., 1984; Itoh et al., 1988; Motzel and Riley, 1992; Franklin et al., 1994). At the same time, inapparent infection is commonly observed and may be inferred from several lines of evidence, chief among them the frequent reports of intercurrent epizootics subsequent to stressful or immunosuppressive treatments, as well as the documented results of immunosuppressive diagnostic screening techniques (Maejima et al., 1965; Takagaki et al., 1967; Karasek, 1970; Yamada et al., 1970; Fries, 1979a; Thunert, 1980; Nakayama et al., 1984; Gibson et al., 1987). By use of antigens derived from infected liver suspensions, it has been established by fluorescent antibody techniques, complement fixation, and agar gel immunodiffusion that the organism is immunogenic and that the antibodies thus induced are protective and usually present in higher titer in younger age groups than in retired breeders (Fujiwara et al., 1965, 1969; Fujiwara, 1967; Onodera and Fujiwara, 1970). Fluorescent antibody techniques, both direct and indirect, have been used to locate C. piliforme antigen in tissues (Fujiwara et al., 1968, 1969; Savage and Lewis, 1972; Fries, 1977a). One technique used to detect inapparence in an earlier era of insensitive serology tests (complement fixation) was the anamnestic response test (ART). This assay was used to detect latent infections (and previous exposure to the agent) in adult rodents that carried antibody levels below the detectable level. The basis of the test is the anamnestic antibody response to a defined challenge with killed bacterial or viral antigens (Fujiwara, 1971). The principle had shown some utility for serologic monitoring of rodent colonies for other pathogens, including mouse hepatitis virus, Sendai virus, C. kutscheri, and Salmonella enteritidis. As a rule, antibody levels were determined on sera from the sixth day after antigen injection. The working assumption had been made that primary antibody responses were detectable only subsequent to the ninth day, whereas those that occur before day 9 are truly anamnestic and reflect sensitization from previous experience with the antigen (Fujiwara, 1971). The assay has fallen into disuse because of the general availability of adequately sensitive immunofluorescent antibody (IFA) and ELISA serologic reagents. Results with immunosuppressive stress tests for confirmation of infection in serologically positive rats have had mixed results and frequently fail to provoke a histopathologically definitive diagnosis. Explanations for failed provocation tests include the following: (1) the detected antibodies were indicators of previous (and transient) infection, i.e., the organism was no longer carried by the animals providing inoculation material for the stress

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test; (2) the particular C. piliforme isolate, although immunogenic, was insufficiently toxigenic to produce lesions; or (3) the antibodies detected were due to antigenically related commensal bacteria and not to C. piliforme. Despite morphologic similarity, isolates from different host species may have substantial biochemical and biologic diversity (Fujiwara et al., 1973a,b; Fries, 1980; Manning, 1993; Riley et al., 1994). The degree of host specificity of C. piliforme isolates and the hazards of interspecific transmission are of great interest and have been studied in some detail. These studies have shown that heterologous transmission (i.e., rat isolate to mouse and vice versa) is less virulent than is homologous transmission, particularly when inoculated by the oral and intravenous routes; that immunity to infective challenge was higher in homologous systems; and that although rat and mouse isolates had certain antigens in common, they also had other unshared antigens (Fujiwara et al., 1971, 1973, 1974; Motzel and Riley, 1991; Franklin et al., 1994). Based on the greater stability of vegetative suspensions from rat liver, Fujiwara et al. (1971) suggested that B. piliformis may originally have been a pathogen of the rat, being forced to mutate to a form with surface changes in heterologous hosts. The general view at present is that regardless of host origin, all isolates must be considered as potentially causative agents of C. piliforme infection and Tyzzer’s disease in heterologous hosts. The sensitivity of B. piliformis to various antibiotics has been investigated primarily from the standpoint of treatment and has shed light on the taxonomic position of the organism by development of antibiograms. These studies have established the effectiveness of cephaloridine, tetracycline, and chloramphenicol for protection against experimental infection (Craigie, 1966a,b; Takagaki et al., 1966; Yokoiyama and Fujiwara, 1971). The emphasis has been on prevention of infection, however, rather than treatment. The importance of good sanitary standards was emphasized previously in the discussion on transmission and factors known to predispose infection. The effectiveness of cage lid filters (microisolators) in preventing infection in enzootic environments has been demonstrated (Boot et al., 1996). The disease may be eradicated and excluded by gnotobiotic principles and maintenance within barriers, although the potential for reinfection of gnotobiotically rederived rat colonies is considerable (Hunter, 1971; Hansen and Mollegaard-Hansen, 1990; Hansen et al., 1992b; Hansen, 1996). Although there is no evidence that C. piliforme may traverse the rat placental barrier in natural infections (to infect gnotobiotic neonates during rederivation), the possibility exists that it may because it has been demonstrated experimentally in immunosuppressed rats (Fries, 1978,1979c) and as a natural occurrence in the guinea pig (Boot and Walvoort, 1984). An additional possibility for

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reinfection of rederived rodent stocks may follow reuse of inadequately sanitized contaminated barriers because of the extreme viability of spores (Hansen and Mollegaard-Hansen, 1990), although based on serology alone these results may be open to question.

G. Corynebacterium kutscheri Infection (Pseudotuberculosis) Pseudotuberculosis in laboratory rats and mice is caused by C. kutscheri, first isolated and described from the mouse in 1894 (Kutscher, 1894). Synonymy with this organism is confusing, both because of the descriptive name for gross lesion morphology (pseudotuberculosis) and because of other similarly named but distinct bacterial entities, for example, Corynebacterium pseudotuberculosis (the PreiszeNocard bacterium) and Yersinia pseudotuberculosis. The synonymy with Y. pseudotuberculosis is additionally confusing because it includes Bacterium pseudotuberculosis rodentium (Schutze, 1928, 1932; Hass, 1938). Both of these latter organisms are culturally and biochemically distinct from C. kutscheri, and there is no evidence that either may cause natural or intercurrent disease syndromes in laboratory rats and mice. Kutscher originally identified the organism as Bacillus pseudotuberculosis murium (Kutscher, 1894). The older literature contains references to Clostridium pseudotuberculosis murium (Reed, 1902; Ford and Joiner, 1968), Corynethrix pseudotuberculosis murium (Schechmeister and Adler, 1953), and Corynebacterium pseudotuberculosis murium (Schechmeister and Adler, 1953), which are regarded as synonyms of C. kutscheri. In addition, as pointed out by Giddens et al. (1968), older references to Clostridium muris (Klein, 1903; Mitchell, 1912) have been cited in reviews on murine pneumonia that describe isolation of Gram-positive diphtheroids from rat pulmonary abscesses consistent with those associated with C. kutscheri.Wilson and Miles (1975)continued with C. murium as the current name for this organism; however, Bergey’s Manual retains the name C. kutscheri. Taxonomically, C. kutscheri is classified in Phylum Actinobacteria, Class Actinobacteria, Order Actinomycetales, Family Corynebacteriaceae, Genus Corynebacterium. The complete genomic sequence of this organism has been determined (Ruckert et al., 2015). Naturally occurring syndromes of clinically apparent disease occur in laboratory rats and mice, and the latent carrier state has been recognized in both of these species as well. The organism has been isolated from wild rats (Boot et al., 1995a) and from humans (Fitter et al., 1979; Messina et al., 1989; Sixl et al., 1989; Holmes and Korman, 2007); thus wild rat carriers represent reservoirs. Similarly, the agent has been isolated from hamsters and guinea pigs, which can act as asymptomatic carriers (Vallee et al., 1969; Nelson, 1973; Amao et al., 1991).

C. kutscheri has been encountered as a primary pathogen in unprovoked infections of the rat (Vallee and Levaditi, 1957; Pestana de Castro et al., 1964; Ford and Joiner, 1968; Giddens et al., 1968; Nelson, 1973; Bhandari, 1976; McEwen and Percy, 1985) as well as the mouse (Bicks, 1957; Bonger, 1901; Hojo, 1939; Kutscher, 1894; Polak, 1944; Savage, 1972; Weisbroth and Scher, 1968a,b; Wolff, 1950). More commonly, however, in both species, inapparent or latent infections have been unmasked in the course of research protocols by experimental treatments lowering host resistance or impairing immunocompetence. Such treatments have included cortisone (Antopol et al., 1951,1953; Berlin et al., 1952; Caren and Rosenberg, 1966; Fauve and Pierce-Chase, 1967; LeMaistre and Thompsett, 1952; Pierce-Chase et al., 1964; Robinson et al., 1968; Speirs, 1956; Yamada et al., 1970), X irradiation (Schechmeister and Adler, 1953; Schechmeister, 1956), and vitamin deficiency (particularly biotin (Gundel et al., 1932; Giddens et al., 1968) and pantothenic acid (Seronde, 1954; Zucker and Zucker, 1954; Seronde et al., 1955, 1956; Zucker et al., 1956; Zucker, 1957)). Some infections with other organisms, e.g., ectromelia virus (Lawrence, 1957) and Salmonella (Topley and Wilson, 1922e1923), have been thought to provoke overt expression of disease, whereas others (e.g., Sendai virus, rat virus, and sialodacryoadenitis virus) have not had that effect (Barthold and Brownstein, 1988). Indeed, the rat was long considered relatively insusceptible. Rats were frequently found refractory to unconditioned transmission attempts (Kutscher, 1894; Bonger, 1901; Seronde, 1954; Ford and Joiner, 1968), and clinical expression was thought to require conditioning by immunosuppressive or nutritionally deficient treatments (Kutscher, 1894; Bonger, 1901; Seronde, 1954). Other modulators of infection are known to include antiinflammatory agents such as L18-MDP, which can restore rat resistance ordinarily ablated by cortisone treatment (Ishihara et al., 1984) and upregulation of galactin-3 (Won et al., 2007). A tumorlytic factor, distinct from TNF-a or -b, has been isolated from the serum of mice injected with T-cell mitogens extracted from C. kutscheri (Kita et al., 1995). Factors stimulating cytokine production, for example, interleukins (IL-1, IL-2, and IL-10) and TNF-a, contribute to nonspecific resistance of rodent hosts (Kita et al., 1992; Jeong et al., 2009, 2012). It is known that heritable resistance and susceptibility occurs among various mouse (Pierce-Chase et al., 1964; Weisbroth and Scher, 1968a; Amao et al., 1993) and rat (Seronde et al., 1955, 1956) strains, and that rat stocks and age groups vary in their capacity for immunologic response to infection (Suzuki et al., 1986, 1988). The concept of latency in the mouse was investigated in some detail by Pierce-Chase and Fauve (Fauve et al.,

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1964, 1966; Pierce-Chase et al., 1964; Fauve and PierceChase, 1967). Their initial observation was that randomly chosen mice from their colony could be placed into one of two categories based on the susceptibility or resistance of the mice to an intravenous challenge with C. kutscheri. They concluded that the resistant mice were actually resistant to superinfection because a state of premunition (i.e., infection immunity) existed in these stocks. The resistant strains, they thought, were immunologically sensitized (and protected) by virtue of preexisting latent infections. They believed the latent organism was an avirulent variant of C. kutscheri carried in the same organs as the virulent form and that shortly after a suitable stress, for example, cortisone administration, these avirulent “A” bacterial populations shifted to the virulent “K” form. These results were not supported by subsequent investigation (Bruce et al., 1969; Hirst and Olds, 1978), and in the light of experience since, “resistance” and “susceptibility” are thought to relate more to (1) host factors of relative immunocompetence and (2) niche limitations to ascendancy imposed by the oropharyngeal and gastrointestinal microbiomes rather than to variants of the agent. It is assumed that the isolation of C. kutscheri from the oral cavity, middle ear, lungs, and (abscessed) preputial glands in the course of routine diagnostic monitoring of essentially normal conventional rats (Matheson et al., 1955; Amao et al., 2002) actually represents cultural confirmation of asymptomatic or latent infection in resistant hosts. In mice it has been shown that C. kutscheri could be culturally detected in the feces of asymptomatic mice for at least 5 months after experimental infection by orogastric gavage (Amao et al., 2008). Presumptive diagnosis may be made at necropsy by smears of affected tissues (or later, in sectioned material), in which typical aggregates of C. kutscheri are demonstrated by Giemsa or Gram stains. Although grayish-blue amorphous bacterial colonies may be seen in tissue sections, typical organisms are not readily discernible in material stained with hematoxylin and eosin. The organisms may easily be demonstrated by Giemsa or tissue Gram (Brown and Brenn or Grame Weigert) stains as irregular palisades or “Chinese letter” arrangements of Gram-positive diphtheritic rods with metachromatic granules. Gram stain reactions of C. kutscheri may be variable in clinical material. Definitive diagnosis requires cultural isolation and biochemical characterization of the organism and may be confirmed by molecular detection of microbial genomic material in tissues or swabs (Saltzgaber-Muller and Stone, 1986; Jeong et al., 2013). Triturates of aseptically collected suspect tissues, swabs, and aspirated washings of the nasal turbinates or middle ears should be cultured directly on 5% blood

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agar plates and incubated aerobically at 37
C. A selective furazolidoneenalidixic acidecolimycin medium may be used to facilitate recovery of low numbers of organisms from nonlesioned or contaminated clinical material (Amao et al., 1990, 1995b, 2008; Brownstein et al., 1985), including feces (Amao et al., 2008). Colonies of C. kutscheri on blood agar are 1e2 mm in size after 24 h, circular, entire, domeshaped, grayish-white in color, smooth, and usually nonhemolytic (whereas C. pseudotuberculosis is hemolytic on blood agar) (Wilson and Miles, 1975). Growth and biochemical characteristics are detailed in the text by Hansen (2000). Isolates of C. kutscheri are facultatively aerobic and nonmotile. Cultures may be propagated on trypticase soy broth or agar. Although C. kutscheri does not form spores, it is quite stable at room temperature in exudates, urine, and feces. The organism is viable in feces held at room temperature without loss of numbers and was still retrievable in culture at 5 days (Amao et al., 2008). It will survive for at least 8 days at 4 and e20
C in phosphate-buffered saline, which was recommended as a transport medium (Shimoda et al., 1991). As a Gram-positive rod, C. kutscheri should be regarded as requiring washing temperatures of at least 180
F to inactivate. The means of spread of C. kutscheri within a colony is not known with certainty, but given that the organism is quite consistently isolated from submaxillary lymph nodes of experimentally infected rats (Brownstein et al., 1985) and from cervical lymph node and oropharynx of subclinically infected conventional rats and mice (Amao et al., 1990, 1995a,b, 2002; Fox et al., 1987), it would seem reasonable to conclude that oral transmission by direct contact or housing on dirty litter are probably the dominant modes. A higher rate of organism recovery has been reported from oral sites than from nasopharyngeal washings (Amao et al., 1995b; Fox et al., 1987), even though rat lungs are the primary site of overt infection. Likewise, given the predilection for kidney abscesses in both host species and high recovery rates from large bowel contents (Amao et al., 1995a), both urine and feces from subclinical carriers should be regarded as infective. Although vertical transmission has been demonstrated experimentally in the mouse (Juhr and Horn, 1975), the marked decline in prevalence of this condition in recent years is testimony to the success of its eradication in gnotobiotically rederived rodents. Because of the chronicity of inapparent infections, the origin of epizootics, when detected, is often obscure. Both morbidity and mortality are extremely variable and depend for expression on the interaction of husbandry, microbiome, heritable factors of resistance or susceptibility, and modulators of immunocompetence (e.g., virus infections that lower resistance). Clinical

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signs in unprovoked rats usually are those associated with subchronic respiratory disease. Variable degrees of upper respiratory signs may be seen, including reddish to mucopurulent discharges from the nares and medial canthus. Affected rats are depressed and anorexic, with humped posture and ruffled haircoat. Respiratory rales are common. Untreated animals typically die 1e7 days after the onset of signs. Any age group may be affected, and a pattern of ongoing but sporadic, low-intensity mortality is typical of enzootically infected colonies. Gross lesions in the rat may be confined to pulmonary tissues and consist of numerous pale foci 1e5 mm in size scattered throughout the parenchyma of the lung. Typically, the lesions are liquefied centrally but may be caseous. Occasional areas of coalescence of foci may be quite large, up to 1.5 cm, and involve substantial portions of a given lobe. The tissue surrounding foci is congested, and entire lobes may be consolidated. Fibrinous adhesions to the thoracic wall are not uncommon and occasionally make retraction of the lung difficult. Such adhesions become fibrous in more chronic cases. Similar foci may less frequently be seen in the liver and kidney of infected rats than in the mouse, in which hematogenous septic embolization and abscessation of organs with capillary nets, e.g., the liver and kidney, are the rule (Weisbroth and Scher, 1968a). Infrequently, arthritic lesions of the pedal extremities and subcuticular abscesses may be seen (Fischi et al., 1931; Nelson, 1973). It was pointed out earlier that abscesses of the preputial glands and middle ears in the rat (via bites from oropharyngeal carriers?) frequently yield C. kutscheri in the absence of pulmonary lesions. Under conditions of cortisone preconditioning (or provocation), the appearance of hematogenous distribution may be more apparent. In considering the issue of how organisms get to the lungs, Giddens et al. (1968) reasoned that inasmuch as the pulmonary lesions of rats were interstitial rather than bronchial, hematogenous embolization to the lung was more consistent with the observations than was inhalation. They also pointed out that pulmonary infection in their own experiments followed oral or intraperitoneal inocula, as did those provoked by the intracardiac injection of C. kutscheri by other investigators (LeMaistre and Thompsett, 1952). Fauve et al. (1964) recovered inapparent C. kutscheri from the lungs of asymptomatic lesionless mice. Microscopic lesions in any of the affected tissues consist of abscesses that vary in size from several micrometers up to the size seen grossly at necropsy. The centers of such lesions are necrotized and, depending on the stage, may be coagulative and relatively acellular early, liquefactive (later) as acute inflammatory cells infiltrate, and mature as inspissated or caseous abscesses. The inflammatory infiltrate surrounding and invading

necrotized areas consists of neutrophils early in the course and includes macrophages, lymphocytes, and plasma cells as the lesions become subacute. Fibroplasia is not pronounced in pulmonary lesions; epitheloid cells, calcification, or giant cells are not seen. Thus although the pulmonic lesions are spoken of in the older literature as being “granulomatous,”Giddens et al. (1968) pointed out that there is little histopathologic support for this description, and the term “pseudotuberculosis” would not really seem to apply. In the lung, suppurative exudates are seen in the bronchioles and bronchi. Interstitial tissues peripheral to pulmonary abscesses are congested, alveoli are filled with proteinaceous fluids, and leukocytes may be seen as perivascular cuffs around blood vessels. Foci are believed to enlarge and coalesce by radial expansion (Weisbroth and Scher, 1968). The use of molecular methods for in situ detection of the agent in latent infections (Saltzgaber-Muller and Stone, 1986) and lesioned tissues (Jeong et al., 2013) has been explored but not widely reported. Both subclinically and overtly infected animals develop antibodies that may be detected by tube or microplate agglutination and indirect immunofluorescence (Brownstein et al., 1985; Suzuki et al., 1986, 1988) or, more effectively, by the ELISA method (Ackerman et al., 1984; Fox et al., 1987; Boot et al., 1995a). Although Weisbroth and Scher (1968b) were unable to detect antibodies in lesionless mice from an enzootically infected colony by tube agglutination, the more sensitive ELISA technology has established the use of this technique in health surveillance programs to serologically detect asymptomatic, inapparently infected rats and mice (Ackerman et al., 1984; Boot et al., 1995a). In enzootically infected breeding colonies, colostral antibodies to C. kutscheri may be detected for the first 3e4 weeks of life, declining to undetectable by the eighth week, rising slowly thereafter to become detectable by the fifth to sixth month and having the highest levels after the eighth month of life (Suzuki et al., 1986, 1988). Although minor differences may exist in homology between isolates from various host species or geographic origin, the great majority of C. kutscheri isolates form a culturally, morphologically, ecologically, and serologically homogeneous group (Yokoiyama et al., 1977; Boot et al., 1995a). Several other methods to detect latent infection have been explored. Cortisone provocation has been extensively used as a means of expressing latent infections and has been advocated for diagnostic surveillance of rat and mouse colonies (Utsumi et al., 1969; Karasek, 1970; Yamada et al., 1970). Similarly, Fujiwara (1971) and his colleagues have explored the use of the ART with this organism and greatly increased the rate of serologic detection of rodents with, at the time, otherwise undetectable asymptomatic infections. Killed

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suspensions of organisms are injected and the test animals bled on the sixth day. Antibody responses, if any, by the sixth day are held to be anamnestic and reflective of previous experience (sensitization) with C. kutscheri. The practical need for both of these tests, that is, immunosuppressive provocation and the ART, has declined in recent years, supplanted by the more practical and costeffective ELISA serology. The use of PCR to detect C. kutscheri in experimentally infected pulmonary tissue has been reported (Jeong et al., 2013). Visibly ill animals can be treated by medication of the water or individual injections of suitable antibiotics after antibiotic sensitivity in vitro has been determined. The organism has been shown to be sensitive in vitro to a variety of narrow- and broad-spectrum antibiotics (Weisbroth and Scher, 1968a), and antibiograms have been developed for C. kutscheri rat isolates (Owens et al., 1975). If the morbidity is low, a number of unprovoked epizootics have been brought under control by the expedient of simply removing visibly ill animals from the colony (Georg, 1960; Weisbroth and Scher, 1968b; Bhandari, 1976). It is emphasized that such procedures, that is, antibiotic treatment and rigid culling, serve to remove the visibly ill tip of the iceberg but leave behind the great bulk of inapparently infected reservoir hosts. The disease may be prevented or excluded from rat colonies by procurement of gnotobiotically derived stock that are housed inside microbiologic barrier systems. This approach has been so successful that there have been no reports of overt C. kutscheri infection in laboratory rats in either naturally occurring outbreaks or those induced by some form of experimental manipulation since the mid-1970s (more than 40 years) with the single exception of a case report in immunosuppressed rats from a conventional facility in Korea (Won et al., 2007). Similarly, to the extent that it may be inferred from the fragmentary and sporadic summaries of rat health surveillance programs (cited in the Introduction to this chapter), neither has the agent been detected asymptomatically in submitted sample cohorts, again with the exceptions occurring in rat samples from conventional facilities in Japan (Amao et al., 2002) and Korea (Won et al., 2006). Thus although it can be concluded that C. kutscheri has been essentially eradicated from gnotobiotically derived stock that are housed inside microbiologic barrier systems the potential for introduction via wild rats or procurement of rats from conventional sources can pose an infectious hazard.

H. Filobacterium rodentium Infection (syn. Cilia-Associated Respiratory Bacillus) Filobacterium rodentium (cilia-associated respiratory bacillus, “CAR bacillus”) infection was originally described as a chronic respiratory disease of aging

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rats, with lesions resembling those of murine mycoplasmosis (Van Zwieten et al., 1980a,b). Clinically, the infection caused signs of respiratory distress and was characterized microscopically by bronchitis, bronchiectasis, mucopurulent exudation, lymphoid follicular hyperplasia, and basophilic striation of airway epithelial cilia. Silver impregnation stains revealed the striations to consist of densely packed bacteria among the cilia bordering the airways. The argyrophilic bacteria were seen from the upper respiratory tract, including the nasoturbinates and middle ears, to the larynx, trachea, and various levels in the lung. The etiological significance of this bacillus was uncertain because the rats involved were concurrently infected with M. pulmonis, Sendai virus, and pneumonia virus of mice. Within several years, similar conditions with morphologically consistent bacterial involvement were reported in a wider range of host species, including Mystromys (MacKenzie et al., 1981), laboratory mice (Griffith et al., 1988) and rabbits (Kurisu et al., 1990), pigs (Nietfeld et al., 1999), and cattle (Hastie et al., 1993; Nietfeld et al., 1999), and additional episodes of infection were seen in both wild (Brogden et al., 1993) and laboratory (Matsushita, 1986; Itoh et al., 1987b) rats. The condition was seen initially in the Netherlands, but within a short period of time reports came from the United States, Italy, Sweden, Spain, Japan, and Australia (Ganaway et al., 1985; Matsushita, 1986; Matsushita et al., 1987; France, 1994; Oros et al., 1997; Caniatti et al., 1998). In many of these earlier reports, there was evidence that other respiratory pathogens were simultaneously present, thus CAR bacillus was relegated to the status of a coinfecting opportunist rather than that of a primary pathogen. This interpretation was supported by PCR analysis of a number of isolates, including that of Ganaway et al., (1985), indicating contamination by Mycoplasma species, including M. pulmonis (Schoeb et al., 1997a). Nonetheless, outbreaks were reported in which careful diagnostic searches could not support concurrent infection by copathogens and in which lesions of chronic respiratory disease could only be attributed to CAR bacillus (Medina et al., 1994). There appears little doubt that CAR bacillus is a primary respiratory pathogen for laboratory rats, mice, and rabbits and as an opportunistic copathogen that may complicate any of a number of rat respiratory diseases, including mycoplasmosis. The name cilia-associated respiratory bacillus and its acronym CAR bacillus were proposed by Ganaway et al. (1985) for use until such time as the organism could be taxonomically classified. Recently, the taxonomic status of CAR bacillus has been established (Ike et al., 2016). The new classification for the organism is F. rodentium gen nov., sp. nov. The type strain is SMR-C. F. rodentium is placed in the Family

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Filobacteriaceae fam. nov. and Order Sphingobacteriales. The 16S rNA gene sequence analysis revealed that SMR-C and other strains of CAR bacillus isolated from rodents all belonged to the Phylum Bacterioides. Strain SMR-C was compared with other CAR bacillus isolates, and found to be deeply nested within the CAR bacillus clade, and it did not cluster with cow, pig, or rabbit isolates of CAR bacillus. CAR bacillus is difficult to cultivate, thus impeding biochemical characterization and taxonomic assignment. Isolation was initially successful in embryonated chickens eggs (Ganaway et al., 1985), then in 3T3 tissue cultures (Cundiff et al., 1994a,b), and finally on cell culture medium supplemented with conditioned medium from tracheal organ culture (Shoji et al., 1992) and unsupplemented artificial media (Schoeb et al., 1993a). F. rodentium is a Gram-negative, argyrophilic, and filamentous rod measuring approximately 0.2  6.0 mm with a triple layer cell wall and bulbous ends. It stains poorly with aniline dyes (and may be missed in tissues stained with hematoxylin and eosin), is not acid fast, is PAS negative, and does not form spores. CAR bacillus is motile but lacks apparent flagella, pili, or axial filaments and has been classified with the “gliding bacteria” (Ganaway et al., 1985). It is heat labile (56
C for 30 min) but survives successive freezing and thawing and remains viable at room temperature for at least 1 week in allantoic fluid preparations (Waggie et al., 1994). Despite morphologic similarity leading to their common nomenclature as CAR bacillus, rat and rabbit isolates are different and somewhat species specific. The rat isolates in culture are more motile, are larger and thicker than rabbit isolates, and form multicell aggregates (Schoeb et al., 1993b; Cundiff et al., 1994a). It has been shown that CAR bacillus isolates differ in their virulence but that there are no important differences in susceptibility to CAR bacillus among Lewis (LEW), Sprague-Dawley,and Fischer 344 (F344) rats (Schoeb et al., 1997b). Mice used for infectivity studies with rat isolates demonstrated colonization of respiratory epithelium, seroconverted, and developed pulmonary lesions, whereas mice inoculated with rabbit isolates may seroconvert but were refractory to infection (Shoji-Darkye et al., 1991; Cundiff et al., 1994a). Rabbits inoculated with rabbit origin CAR bacillus developed nasal discharge and microscopic lesions of the respiratory tract and seroconverted, but they were not colonized and did not seroconvert when inoculated with rat origin CAR bacillus (Matsushita et al., 1989; Cundiff et al., 1995a). Sequence analysis of the 16S rRNA gene from rabbit and rat isolates indicated these isolates were quite different (with only 48.8% homology), suggesting they probably should be classified in different genera. Rabbit isolates identified most closely with Helicobacter (about

90% homology), and it was concluded the isolates were host specific and belonged to separate genera (Cundiff et al., 1995a). The genetic and antigenic relationships remain quite complicated because all isolates share certain epitopes (Hook et al., 1998). Analysis of rat isolate 16S rRNA sequences indicated that rat origin CAR bacillus is most closely related to Flavobacterium and Flexibacter (Schoeb et al., 1993b; Kawano et al., 2000). Because it has been shown experimentally that transmission can occur between rodent species, it must be assumed that rodent carriers of CAR bacillus may be infectious for other rodent species (Shoji et al., 1988b; Matsushita et al., 1989). Infection appears to be most effectively transmitted by direct contact (Matsushita et al., 1989) and, in enzootically infected breeding colonies, may be transmitted by dams to neonates as early as 1e2 weeks after birth (Ganaway et al., 1985). Dirty bedding from infected animals fails to transmit the infection to exposed sentinels (Cundiff et al., 1995b). The clinical signs and corresponding pathology of CAR bacillus infection are quite variable, and expression depends on the contending interplay of modulators such as isolate qualities, host age, degree of heritable resistance, and the effects of resistancelowering copathogens. Infected rat carriers may be asymptomatic (Schoeb et al., 1997a; Shoji et al., 1988a; Cundiff and Besch-Williford, 1992). Nonspecific signs of respiratory diseasedincluding labored breathing, persistent rales, nasal discharge, periocular porphyria, head tilt, hunched posture, roughened haircoat, and weight lossdhave been observed (Matsushita and Joshima, 1989; NRC, 1991; Cundiff and Besch-Williford, 1992). Similarly, the gross lesions of CAR bacillus infection seen at necropsy are not distinctive. They can include the presence of exudates in the nasoturbinates and middle ears (tympanic bulla) and serous to mucopurulent exudates in the trachea. The lungs may fail to deflate because of mucopurulent plugging, and there may be areas of consolidation or scattered foci on the pleural surface. These lesions, of course, are quite similar to those of murine respiratory mycoplasmosis, in the past a frequent copathogen, and the question is open as to how much of the pathology ascribed prior to 1970 to mycoplasmosis (alone) may really have been caused by complicated infections with both agents. Experiments designed to ensure the absence of M. pulmonis and other possible copathogens have clarified and confirmed the ability of CAR bacillus to cause the lesions of chronic respiratory disease (Ganaway et al., 1985; Ganaway, 1986; Matsushita and Joshima, 1989). These studies have shown that CAR bacillus infections initiate as an upper respiratory condition. The earliest lesions (rhinitis, tympanitis, and tracheitis) of experimental infection in rats occur in the ciliated epithelium of the nasoturbinates, eustachian tube, middle ears,

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and trachea. Microscopically, chronic inflammatory infiltrates are seen in the lamina propria underlying the ciliated epithelium, and with silver stains, argyrophilic, filamentous rod-shaped bacteria can be observed on the surface of the ciliated border. Later, the infection descends to the pulmonary tree and microscopic lesions of mucopurulent bronchitis and bronchiolitis develop, along with prominent peribronchial cuffs of chronic inflammatory cells (Itoh et al., 1987b). Ulcerative bronchiolitis and mucopurulent alveolitis occur in natural CAR bacillus disease, but are not typically observed in M. pulmonis infections (Schoeb et al., 1997a). The ciliated epithelium lining the airways is markedly infiltrated with chronic inflammatory cells, predominantly lymphocytes and plasma cells. The apical insertions of CAR bacilli have been demonstrated by electron microscopy (see Van Zwieten et al., 1980a,b). Excellent light and electron microscopy photomicrographs of CAR bacillus may also be seen in the work by Ganaway et al. (1985). Health surveillance screening tests to detect CAR bacillus infections in rats are principally oriented to serologic tests for antibodies indicative of exposure and PCR tests to detect genomic material within the rat respiratory system. Both immunofluorescence assay (Matsushita et al., 1987) and ELISA (Ganaway et al., 1985; Shoji et al., 1988a; Thuis and Boot, 1999) are commonly used as serologic screening tests. Although the potential exists for cross-reactive false positives because of shared antigenic epitopes with other bacterial forms (Hook et al., 1998), this has not proven to be a practical limitation to the use of serodiagnostics. The optimum age for serologic detection of rats from infected breeding colonies is 10e 14 weeks of age, based on the sequence that rat pups born of antibody-positive dams have colostral titers for the first 3e4 weeks of life, which will wane to become low or negative by 5e7 weeks (Ganaway et al., 1985; Matsushita et al., 1987; Shoji et al., 1988a). Screening tests for detection of CAR bacillus infection also include PCR and reverse transcriptionPCR, which offer means for overcoming some of the disadvantages of serology, for example, the potential for false positives and the problem that rats sampled for testing, although exposed and colonized, may not yet have developed maximal titers (Cundiff et al., 1994b; Goto et al., 1995). Antemortem (or necropsy) samples for PCR screening tests may be taken by swab with a higher rate of recovery from the nasoturbinates (Franklin et al., 1999) than that of samples from the oral cavity (Goto et al., 1995). Diagnostic confirmation of screening testing that is positive from either asymptomatic or clinically affected rats requires demonstration of the organism in infected tissues. Confirmation may be obtained by light microscopy with special stains, by immunofluorescent or immunoperoxidase methods to demonstrate CAR bacillus in tissues (Oros et al., 1996), by isolation of the

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organism in culture, or by some combination of these approaches. One caveat here is that resistant rodent strains may only have brief periods of a few weeks when the organism is in residence (Shoji-Darkye et al., 1991), thus older rats may be serologically positive but no longer infected. Because it has been shown that upper respiratory involvement precedes pulmonary involvement (Itoh et al., 1987a,b), necropsy tissue selection for diagnosis of CAR bacillus infection should include tracheal sections or smears. An improved method for demonstrating the organism in tracheal scrapings has been reported (Medina et al., 1996). Transmission of the organism within contaminated facilities is principally by direct contact. Although sentinels in contact with dirty bedding fail to contract CAR bacillus infection (Cundiff et al., 1995b), fomites may be involved in the spread of the organism. Other sentinel studies have indicated probable aerosol transmission within rooms of infected rats (Medina et al., 1994). Ordinary quarantine procedures should be regarded as sufficient to contain the infection to contaminated rooms in a facility. A report on the prevalence of infectious agents in laboratory mice and rats (Pritchett-Corninget al., 2014) indicated a CAR bacillus prevalence in rats of 0.27% (North America) and a 4.63% prevalence in Europe from 26,000 serologic tests. In mice, the combined North American and European prevalence was 0.01% from over 158,000 serologic tests. It should be noted that this report was based on testing rats and mice primarily from pharmaceutical and biotech companies rather than academia.

I. Salmonella sp. Infection (Salmonellosis) Salmonella infections were among the first to be recognized as naturally occurring enzootic or epizootic diseases of laboratory rats and mice. Indeed, the first isolation of Salmonella typhimurium was made by Loeffler in the course of an epizootic of laboratory mice in 1892 (Pappenheimer and von Wedel, 1914). The scientific literature of the next three decades (1895e1925) was particularly rich on the subject of rat salmonellosis and dealt with perhaps three major themes: incidence and significance of salmonellosis in wild rats as vectors of human food contamination, the use of “rat viruses” (Salmonella cultures) for biological control of wild rat populations, and the documentation and description of intercurrent salmonellosis in laboratory rat colonies. The concept of using Salmonella cultures as rodenticides was first suggested in 1893 by the success observed by Loeffler with the use of S. typhimurium in combating a plague of field mice in Thessaly (Pappenheimer and von Wedel, 1914). Another of the early isolates was the Clostridium (S. enteritidis) isolated by Danysz (1914) from an epizootic of field mice in Charny-en-Seine, France. The isolate was observed to have considerable

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pathogenicity for rats. These agents were known as rat viruses and used as popular rodenticides for about 15 years (1895e1910). Such cultures as the Liverpool virus, the Azoa virus of Wherry, the Danysz and Issatschenko viruses, and Ratin I and II (all isolates of S. enteritidis) were widely used on the European continent, England, and in the United States as rat “poisons” (Bahr, 1905; Mallory and Ordway, 1909; Bahr et al., 1910; Pappenheimer and von Wedel, 1914; Jordan, 1925). They were used later for this purpose in Eastern Europe (Trakhanov and Kadirov, 1972). The predictable curtailment of this usage came about when it was recognized that the Salmonellas could not be safely contained in the field, and they were, in fact, responsible for numerous human epidemics (Savage and White, 1923; Jordan, 1925; Bartram et al., 1940). Nomenclature for Salmonella is summarized in Table 12.2. The genus Salmonella contains two species: Salmonella enterica and Salmonella bongori (Fookes et al., 2011; Timme et al., 2013). Whereas S. enterica infects warm-blooded animals and humans, S. bongori is predominantly a pathogen of cold-blooded animals and only rarely has been associated with human infections. S. enterica is divided into six subspecies: enterica, salame, arizonae, diarizonae, houtenae, and indica. These subspecies are further subdivided into >2500 serovars (serotypes). Serovars causing disease in humans and other warm-blooded animals principally belong to S. enterica subspecies enterica. S. enterica subspecies enterica has numerous serovars, including typhimurium and enteritidis (which are not italicized because serovar [serotype] designations have no standing in nomenclature), and are frequently abbreviated for convenience as S. Typhimurium and S. Enteritidis, respectively. Both have been recovered with sufficient frequency to be regarded as typical for murine salmonellosis. The older literature contains a confusing synonymic array for both serotypes, as would be expected for a large bacterial group undergoing taxonomic study for almost 100 years. The synonyms for S. Typhimurium include Clostridium typhimurium, Bacterium aertrycke, S. pestis caviae, S. aertrycke, and S. psittacosis. In like manner, the following are regarded as synonymous with S. enteritidis: Bacterium enteritidis, Clostridium enteritidis, Danysz Clostridium, S. enteritidis var. danysz, and Gaertner’s (or Ga¨rtner’s) Clostridium. Both S. Typhimurium and S. Enteritidis have essentially identical epizootiology in rats, and the discussion that follows applies to both, except where noted. Salmonellas isolated from both wild and laboratory rats before the 1950s most commonly included S. Typhimurium and S. Enteritidis. Although many rodent infections with these agents are subclinical, they are quite pathogenic for rats (and mice) and the following pathologic descriptions are based on infections caused by these two serotypes.

As mentioned previously, surveys of wild rat populations have established that these animals were, and remain, enzootic vectors of human Salmonella food poisoning, particularly of dairy, bakery, and meat products (MacAlister and Brooks, 1914; Savage and White, 1923; Salthe and Krumweide, 1924; Jordan, 1925; Meyer and Matsumura, 1927; Verder, 1927; Kerrin, 1928; Khalil, 1934; Nafiz, 1935; Staff and Grover, 1936; Bartram et al., 1940; Welch et al., 1941). Although the incidence of survey reports has declined in recent decades, such studies indicate the continuing association of wild rats with Salmonella (Varela et al., 1948; Lee, 1955; Brown and Parker, 1957; Bruce et al., 1969; McKeil et al., 1970). It should be noted that in addition to Salmonella food poisoning, zoonotic infections from pet rodents remain as a public health concern (Swanson et al., 2007). Similarly, wild rats must still be regarded as significant hazards for the introduction of salmonellas to laboratory rat colonies. A wild rat source was shown to be the cause of a waterborne contamination that survived (faulty) chlorination in one of the few reported episodes of salmonellosis in laboratory rats since 1939 (Steffen and Wagner, 1983). S. Typhimurium and S. Enteritidis are still occasionally encountered in laboratory rodents (usually with a wild rodent source in the history), but in recent years, most salmonellas isolated from laboratory animals are species more suggestive of originating from some other source. These sources include dietary components, drinking water, bedding materials, and avian or human contacts. The Salmonella serotypes encountered include the following: Agona, Amsterdam, Anatum, Binza, Blockley Bredeney, California Cerro, Infantis, Kentucky, Livingstone, Montevideo Oranienburg, Poona, Senftenberg, Tennessee, Mbamdaka, and others (Simmons and Simpson, 1980, 1981; Kirchner et al., 1982; Lentsch et al., 1983; Seps et al., 1987). These ordinarily nonrodent salmonellas tend to be less pathogenic for rats and mice and typically occur as essentially subclinical infections. Sporadic but quite current isolation of these nonrodent serotypes underscores the continuing importance of maintaining the high sanitary standards associated with rodent barrier facilities, including treatment of diets, beddings, and drinking water and personnel hygiene practices that include use of protective outerwear. Salmonella infections had been internationally regarded as among the most important of laboratory animal diseases. The renown was not misplaced since it was a serious and common disease of laboratory rats for about four decades (1890e1930), as the frequent reports during that period indicate (Bainbridge and Boycott, 1909; Boycott, 1911; Pappenheimer and von Wedel, 1914; Cannon, 1920; Ball and Price-Jones, 1926; Price-Jones, 1926e1927; Friesleben, 1927; Bloomfield and Lew, 1943a,b,c). Much of the earlier literature, before

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1914, has been summarized by Pappenheimer and von Wedel (1914). The naturally occurring disease in rats has been reported on several occasions subsequently (Buchbinder et al., 1935; Jones and Stewart, 1939; Ratcliffe, 1949; Taylor and Atkinson, 1956; Sacquet, 1959; Kunze and Marwitz, 1967; Potel, 1967; Ray and Mallick, 1970; Lentsch and Wagner, 1980). The decreasing prevalence of this condition is worth emphasizing because it has only been reported in laboratory rats in the United States once (Lentsch and Wagner, 1980) since 1939. The epizootiology of this disease has undergone a radical transition in American rodent stocks, attributable perhaps in equal measure to the concepts of gnotobiotic technology with the principle of disease exclusion, to the elevation of standards of animal care generally, and as a consequence of US federal laws (namely, 91:579, the Animal Welfare Act and amendments) and the accreditation guidelines of the Association for Assessment and Accreditation of Laboratory Animal Care. In most countries with similar standards, the disease is comparatively rare. The clinical form of the disease in rats may vary from that of an acute epizootic with substantial mortality to that of a clinically latent disease that may be silent until the animals are placed under investigatory stress. Many factors are known to modulate the clinical expression of Salmonella infections in laboratory rats and mice. They include the level of acquired immunity, genetic resistance, virulence and inoculum concentration of the organism, age of the host, nutritional balance, and any of a variety of stressors that may act to lessen the immunologic competence or general vitality of the host (Topley and Wilson, 1922e1923; Webster, 1924; Guggenheim and Buechler, 1947; Bohme et al., 1954; Habermann and Williams, 1958; Blanden et al., 1966; Mackaness et al., 1966; McGuire et al., 1968; Newbeme et al., 1968; Kampelmacher et al., 1969; Groschel, 1970; Robson and Vas, 1972; Tannock and Smith, 1972; Marecki et al., 1975). It is also pointed out that the pathologic consequences of Salmonella infection as a monocontaminant of germ-free rats and mice (Margard and Peters, 1964; Wostman, 1970; Ruitenberg et al., 1971) are considerably attenuated in comparison to microbially associated animals, thus the floral constitution of the host (and the issue of colonization resistance) comes into play as well. After the acute phase, natural infections in laboratory rats from enzootically infected colonies tend to moderate thereafter with little subsequent clinical signs or mortality. Clinical signs of salmonellosis include reduced activity, thirst, ruffled fur, hunched posture, anorexia, weight loss, and softer lighter stools (Pappenheimer and von Wedel, 1914; Buchbinder et al., 1935; Ratcliffe, 1949; Maenza et al., 1970). Diarrhea has been an inconstant sign under conditions of natural infection, seen more

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commonly during acute episodes, and less frequently in enzootically infected colonies (Buchbinder et al., 1935). Experimental infections of rats with S.Typhimurium by Maenza et al. (1970) have established the brief incubation period of 2e3 days before development of clinical signs, including diarrhea. Diarrhea incidence averaged 15%e20% of infected rats but was increased in rats that fasted before infection and by overcrowding. Maenza et al. confirmed the results of others that the diarrheal sign was dose dependent (Maenza et al., 1970). The incidence of diarrhea was maximal 4 days after infection and ranged in persistence from 1 to 21 days. The character of the stools in rats with diarrhea ranged from formless but normally colored, to mucoid and occasionally bloody bowel movements. Experimental Salmonella diarrhea has been extensively investigated as a diarrhea model in rats (Wostman, 1970; Powell et al., 1971a,b). The gross findings in fulminating and acute cases are limited to those consistent with septicemia. Most abdominal organs, particularly the liver and spleen, are congested, and lesions may be thus limited. Subchronic and chronic cases, with longer duration before death, generally have specific lesions. The disease is preeminently a classic enteric infection of the ileum and cecum; however, intestinal lesions may extend on either side of this location. Bacteremic or septicemic ascension to the liver and spleen occurs commonly. It is doubtful if the cardiac and pulmonary lesions described in the older literature (Pappenheimer and von Wedel, 1914) are Salmonella related, because they have not been described by more recent investigators (Buchbinder et al., 1935; Naughton et al., 1996). Gross lesions of the intestines include thickening of the ileum and cecum with intraluminal accumulations of semiliquid feces, gas, and flecks of blood. The mesenteric and serosal vessels are congested, and the mesenteries edematous (Buchbinder et al., 1935). In animals with diarrhea, the intestinal lesions typically extend into the jejunum and include as key features large and small ulcers of the cecal epithelium and, less commonly, the terminal ileum. The ulcers are usually covered with fibrinous exudates. Microscopically, the ileal and cecal lesions are seen to consist of diffuse enteritis with infiltration of the lamina propria with macrophages, neutrophils, and, later, lymphocytes. The crypt epithelium is typically hyperplastic; however, the villous epithelium shows degenerative changes. Ulcers of the ileal mucosa, infrequently present, tend to be microscopic in size, whereas those of the cecum are larger (up to 20 mm with irregular outline) and grossly apparent. Cecal ulcers were not always accepted as occurring as direct consequences of salmonellosis in rats, and the relationship had been seriously questioned (Jones and Stewart, 1939; Stewart and Jones, 1941; Bloomfield and Lew,

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1943a,b,c, 1948; Bloomfield et al., 1949). They are now recognized, however, as lesions attributable to Salmonella infection (Buchbinder et al., 1935; Maenza et al., 1970). It must be emphasized that subclinically infected animals without diarrhea have similar but less extensive lesions on both gross and microscopic levels of observation. A positive correlation has been shown to exist between the signs of diarrhea and the severity of ileal lesions. Similarly, the persistence of diarrhea is related to the severity of ileal lesions, with sign subsiding as the tissues heal. The intestines of surviving animals were histologically healed by 4e5 weeks after infection (Maenza et al., 1970). Grossly apparent indications of reticuloendothelial involvement are initiated early in the course and include enlargement of the Peyer’s patches of the ileum, the mesenteric lymph nodes, and spleen. Microscopically, the Peyer’s patches and lymph nodes are edematous, have inflammatory infiltrates, and may show focal necrosis or granuloma formation. Splenomegaly is a common feature (Thygessen et al., 2000). Such spleens are dark red or have a grayish capsule and may approximate 4 cm in length by 1 cm width (Pappenheimer and von Wedel, 1914; Buchbinder et al., 1935). Microscopically, the splenomegaly is seen to be caused by intense congestion of splenic sinuses, with spilling over of erythrocytes into the white pulp. Extensive inflammatory infiltrates, particularly of phagocytic cells, are seen in splenic tissues. Focal lesions varying from necrosis to granulomata are commonly seen. The liver is variably congested and dark red or pale and friable. Focal lesions (1e2 mm) may be seen grossly under the capsule and on cut surface but more commonly are not. On microscopic examination, however, small areas of coagulative, acellular necrosis are seen, as has been described in mice (Tuffery and Innes, 1963). Granuloma formation in the rat liver has been described (MacAlister and Brooks, 1914). In the original histopathologic description provided by Pappenheimer and von Wedel (1914), small microthrombi (cell fragment thrombi)dcomposed of mononuclear phagocytes containing cell fragments and bacteria, degenerating phagocytes, cellular debris, and fibrindwere observed in organs with focal necrosis, for example, spleen, liver, and lymph nodes. The investigators concluded that the focal necroses were actually sequelae of embolically occluded (thrombosed) microvasculature. One is tempted to recall, in support of this concept, the pulmonary thrombi in hamster salmonellosis described as pathognomonic by Innes et al. (1956). Certain lesions seen experimentally by using the rat Salmonella modeld for example, cholangitis (Arora and Sangal, 1973), placentitis (Hall, 1974), and polyarthritis (Volkman and Collins, 1973)dhave not been observed in natural infections.

Diagnosis of salmonellosis in the rat is based on a consistent case history, isolation of the organism in pure culture, and cultural and immunologic confirmation of the identified isolate as one of the Salmonella serotypes. It has been pointed out, in the mouse, that difficulty should not be experienced in the retrieval of Salmonella species encountered during acute epizootics (Morello et al., 1964). Large numbers of animals die of the infection, and the many ill animals shed high ratios of Salmonella organisms into the feces. More problems have been encountered in the diagnosis of salmonellosis in asymptomatic carrier animals from enzootically infected colonies. The asymptomatic carrier animal (mouse) has been investigated in some detail (Jenkin et al., 1964). It is understood from such studies that the relationship with the host is one of intracellular multiplication by the organism within a milieu of humoral antibody. Exacerbation of the disease in recovered carriers is prevented by specific antibody, but complete elimination is inhibited by smoldering replication of the bacteria within macrophages. Such animals may shed Salmonella intermittently in the feces over long periods of time and represent infective reservoirs of the disease. Difficulty of detection because of episodic shedding is compounded by the problem that the prevalence of asymptomatic carriers may range from a high 50% of the animals to as low as 5%, thus introducing a substantial risk of fecal sampling error (NRC, 1991). That essentially the same factors are involved in the rat is generally accepted, and the asymptomatic carrier rat has been amply documented (Buchbinder et al., 1935; Naughton et al., 1996). Although clearly of great diagnostic importance, the focus of infection in recovered carriers is not well understood and probably is not the same in each carrier. Several studies have attempted to localize the most likely organs or other locations in carrier animals for more efficient diagnostic retrieval (Buchbinder et al., 1935; Slanetz, 1948; Hoag and Rogers, 1961; Margard et al., 1963; Hoag et al., 1964; Morello et al., 1964, 1965). The consensus is that although a greater number of isolates may be retrieved from intestinesdcompared with liver, spleen, heart, or other organsdnone of these tissue locations results in higher percentage of isolations than from feces alone. For this reason, diagnostic monitoring programs for murine salmonellosis have been oriented to fecal culture. Salmonella is quite stable in dried feces and on environmental surfaces. Strains inoculated in media can survive at room temperature (23
C) for more than 1 year (Kagiyama and Wagner, 1994). As a Gram-negative rod, it is quite sensitive to heat and is killed by exposure to temperatures of 55
C for 1 h or of 63
C (145
F) for 20 min. Note that the pasteurizing temperatures at which rodent diets are pelleted are sufficiently borderline in terms of time and temperature to variably inactivate

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Salmonella (NRC, 1991). Cage washing machine temperatures of 82
C (180
F) are sufficient to inactivate Salmonella. Ordinary disinfectants, including sodium hypochlorite, iodophore detergents, ethanol, phenol, and formaldehyde, are effective (Kagiyama and Wagner, 1994). The factors relating to efficacy of retrieval of Salmonella from rat and mouse feces under varying conditions have been studied. It is known that recovery from individual animal samples is higher than that from pooled samples (Slanetz, 1948; Margard et al., 1963), that the highest number of isolates will be obtained from the first six fecal pellets in any given animal (Margard et al., 1963), and that storage up to 8 days in fluid media does not result in greater degree of isolations than dry feces stored for 8 days (Margard et al., 1963). Similarly, storage temperatures (whether frozen, at room temperature [20
C] or at 40
C) did not affect the percentage of isolations (Margard et al., 1963). It is recommended that feces shipped for diagnostic culture be kept cool and dry to minimize growth of species other than Salmonella that may interfere with isolation. In like manner, primary culture media used for isolation of the organism before biochemical characterization have been explored in great detail. The consensus is that the highest numbers of isolations follow the sequence of overnight enrichment in a broth medium to permit preferential multiplication of Salmonella (compared with coliforms) followed by plating on differential solid agar media. The media have been thoroughly reviewed by Margard et al. (1963), but they and others (Haberman and Williams, 1958; Morello et al., 1964) recommend the sequence of incubating fecal pellets or tissue macerates in selenite-F plus cystine broth overnight followed by streaking onto brilliant green agar as the method of choice. Suspicious colonies are picked from the brilliant green plates and inoculated on triple sugar iron. Isolates consistent with Salmonella sp. are then confirmed by agglutination in polyvalent antisera and characterized biochemically according to their profile (Hansen, 2000). The serotyping of isolates is beyond the competence of most laboratories, and characterized isolates should be forwarded to a recognized diagnostic center for complete identification. Rapid screening and diagnosis of clinical episodes has been described using selective media (e.g., selenite and Hektoen agar) for isolation and the MALDI-TOF MS procedure for identification (Dieckmann and Malony, 2011; Sparbier and Weller, 2012). Both rats and mice respond immunologically to infective challenge by Salmonella (Boycott, 1919; Topley and Ayrton, 1924; Ball and Price-Jones, 1926; Buchbinder et al., 1935; Slanetz, 1948; Hobson, 1957; Kent et al., 1976; Collins, 1968, 1970). Traditionally, it has been

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thought that postrecovery antibody levels, especially in immune asymptomatic carriers, are too low and inconsistent to be of great value in diagnosis. Numerous studies have documented the two-to threefold higher number of successful isolations of Salmonella from feces compared with detection of antibodies in the same animals (Slanetz, 1948; Hobson, 1957; Sacquet, 1958, 1959). For this reason, immunodiagnosis has not been generally recommended, other than as an adjunct to cultural methods for diagnosis. Of the methods explored, complement fixation was found to be more sensitive than was tube agglutination (Morello et al., 1964). Application of ELISA methodology appears to offer promise as a means of detecting low postrecovery antibody levels in carriers found negative by less sensitive serologic tests (Lentsch et al., 1981). Salmonella can be detected by PCR and may detect as few as 10e50 bacterial cells in the sample (Soumet et al., 1994; Stone et al., 1995). Vaccination has not been endorsed as a method of prevention, although the concept has been seriously explored. Vaccination with bacterins or bacteriophage has not been completely effective in either preventing infection in vaccinated individuals or in preventing spread when vaccinated and nonvaccinated mice were placed in close proximity (Topley and Wilson, 1922e1923, 1925; Topley et al., 1925; Ibraham and Schutze, 1928; Walker et al., 1964; Badakhsh and Herzberg, 1969; Rauss and Kalovich, 1971; Cameron and Fuls, 1974). The concept of treatment of infected individuals has been discouraged because of the incomplete effectiveness of various treatment regimens (Habermann and Williams, 1958). The problem has been that although the majority of animals may be apparently freed of Salmonella, a residue of infection always remains to act as a reservoir for rebound of infection in susceptible animals (Bruner and Moran, 1949). Similarly, although fecal testing programs oriented to detection and culling of infected animals may seem to be statistically effective, it is almost impossible, with certainty, to eradicate the infection from a given colony by these means (Steffen and Wagner, 1983). Current recommendations favor destruction of colonies in which the disease has been detected, followed by replacement with gnotobiotically derived stock known to be free of Salmonella. There is no evidence that the organism may traverse the placental barrier.

J. Pasteurellaceae Infection As a group, the Pasteurellaceae are Gram-negative, fermentative, nonmotile coccobacilli that are oxidase and catalase positive and that variably fail to grow on MacConkey agar. Pasteurellaceae are taxonomically complex and as more precise methods have been

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developed and studied, nomenclature of genera and species have evolved. The three classic genera have been Pasteurella (1897), Actinobacillus (1910), and Haemophilus (1917). Within the past decade, the taxa of the rodent Pasteurellaceae have been very significantly revised through genetic and phenotypic studies. Rodent isolates classified as Pasteurellaceae have been reclassified into six new genera: Rodentibacter (Adhikary, S. et al., 2017b), Muribacter (Nicklas et al., 2015), Cricetibacter, Mesocricetibacter (Christensen et al., 2014), Mannheimia (Christensen et al., 2011a,b), and Necropsobacter. These newly reclassified Pasteurellaceae, their previous taxa, along with their primary and additional hosts, are summarized by Adhikary et al.(2017a,b) and Boot et al. (2018). These genera and species are defined as “validly published” by the International Code of Nomenclature of Bacteria, and subsequently were published in the International Journal of Systematic and Evolutionary Microbiology as original articles or in Validation Lists. Accordingly, prior nomenclature for rodent Pasteurellaceae should no longer be used. This new classification now provides a firm basis for the correct phenotypic identification of rodent Pasteurellaceae and delineating strains for pathogenicity studies (Boot et al., 2018). P. pneumotropica is the most commonly recognized Pasteurellaceae associated with mouse and rat infections, and until recently had been thought to be a single species with two biotypes: Jawetz and Heyl. These two biotypes are now reclassified within the genus Rodentibacter as two distinct species: Rodentibacter pneumotropicus and Rodentibacter heylii. R. pneumotropicus is the type species for the genus. In the past, few reports in the literature specified the Pasteurella biotype, Jawetz or Heyl, involved in the study. Therefore lacking identification of the biotypes for reference in these studies, P. pneumotropica will be used in this section rather than the newly, validly published R. pneumotropicus and R. heylii species (Boot et al., 2018). As a health monitoring strategy, FELASA (Mahler et al., 2014) recommends monitoring for P. pneumotropica at 3-month intervals, and for other Pasteurellaceae, screening can be conducted according to the needs deemed necessary by the institution. The report acknowledges that the inclusion of the Pasteurellaceae family is controversial due to the difficulty of some commercial kits to correctly identify P. pneumotropica, as well as the fluidity of the correct phenotypic classification. 1. Rodentibacter pneumotropicus, Rodentibacter heylii (syn. Pasteurella pneumotropica) The most frequently reported rodent pathogen of the Pasteurellaceae is P. pneumotropica. However, when examined critically, it should be recognized that in

many of these reports, the phenotypic characteristics presented do not allow definitive identification to the now recognized species level (see Mutters et al., 1989 for cultural criteria). The Heyl and Jawetz biotypes are genetically distinct (less than 30% related by DNAe DNA hybridization), and both biotypes appear related more to the genus Actinobacillus than to Pasteurella (Ryll et al., 1991; Bisgaard, 1995). On the basis of cultural criteria, DNAeDNA hybridization, and randomly amplified polymorphic DNA analysis by PCR, biotype Heyl isolates are genetically homogeneous, but biotype Jawetz isolates form a genus-like cluster that even contains some V-factor (NAD)-dependent strains (Ryll et al., 1991; Weigler et al., 1996; Schultz et al., 1977; Kodjo et al., 1999). Since its original description (Jawetz, 1948), P. pneumotropica has been associated mainly with the respiratory tract of laboratory rodents, a predilection emphasized in the species name (Jawetz, 1950; Jawetz and Baker, 1950). Sporadic reports of pasteurellosis in rats occurred before that time, notably those of Schipper (1947) and Meyer and Batchelder (1926). Similarly, the older literature contains references to respiratory infection in rats with organisms that on close examination resemble what are now recognized as Pasteurellaceae (Hoskins and Stout, 1920; Matheson et al., 1955; Harr et al., 1969). In these reports, the role of viral and other bacterial agents is undefined and may have been copathogens contributing to the respiratory outbreaks reported. P. pneumotropica has emerged since the 1950s as an important and globally distributed infectious agent of laboratory rats. A report based upon data from a major commercial rodent diagnostic laboratory found that of 8241 cultures sampled, the prevalence of P. pneumotropica was 4.81% (Pritchett-Corning et al., 2009). The majority of samples were from pharmaceutical and biotech companies with the remainder from academic and governmental institutions in North America and Europe. The authors clarify that the reported prevalence is based upon samples and not animals, and that sampling of more than one organ/tissue would have increased the reported ratprevalence figures. Also noted was that the isolates were identified using API 20NE strips, and that a minority of colonies identified as P. pneumotropica by biochemical methods were negative by PCR for the two biotypes, Jawetz and Helyl. In the same paper, there was a reported prevalence in mice of 12.9% from 109,400 samples. Microbiological testing of rats from 161 universities and 101 pharmaceutical firms in Japan found a P. pneumotropicaprevalence of 4.35% and 0.99%, respectively (Hayashimoto et al., 2013). It has been described as a clinical entity in laboratory mice on a number of occasions (Jawetz, 1950; Gray and

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Campbell, 1953; Heyl, 1963; Brennan et al., 1965; Wheater, 1967; Weisbroth et al., 1969; Black, 1975; Needham and Cooper, 1976; Wilson, 1976; McGinn et al., 1992; Grossman et al., 1993; Dickie et al., 1996; Marcotte et al., 1996; Artwohl et al., 2000; Macy et al., 2000; Adhikary et al., 2017a,b). P. pneumotropica may be carried asymptomatically in hamsters (Brennan et al., 1965; McKenna et al., 1970; Sparrow, 1976; Lesher et al., 1985), guinea pigs (Sparrow, 1976; Boot et al., 1995b), and rabbits (Sparrow, 1976). P. pneumotropica is commonly isolated from wild rats and mice (Van der Schaaf et al., 1970; Boot et al., 1986). These other host species should be regarded as reservoirs of potential P. pneumotropica infectivity for laboratory rats. Although encountered less commonly in recent years, a number of earlier surveys have reported P. pneumotropica in breeding stocks of rats and mice (Casillo and Blackmore, 1972; Hill, 1974; Hooper and Sebesteny, 1974; Rehbinder and Tschappat, 1974; Young and Hill, 1974; Sparrow, 1976), frequently in the absence of clinically overt disease. Indeed, inapparent infection has remained one of the characteristics of the organism. Even under conditions of experimental infection, the organism may be clinically silent (Van der Schaaf et al., 1970; Jakab and Dick, 1973; Hill, 1974; Burek et al., 1972). Alternatively, P. pneumotropica may occupy the role of secondary opportunist or copathogen, synergistically enhancing the pathogenicity of a number of respiratory agents, including Sendai virus (Jakab and Dick, 1973; Jakab, 1974), M. pulmonis (Lutsky and Organick, 1966; Brennan et al., 1969a; Eamens, 1984), and P. carinii (Macy et al., 2000). Finally, it may appear very infrequently as a primary pathogen (Brennan et al., 1969a,b; Van der Schaaf et al., 1970). However, possible or known coinfection with viruses or M. pulmonis in these early reports clouds the status of P. pneumotropica as a primary pathogen in the respiratory tract. Little is known about the triggering mechanisms promoting progression from inapparence to pathogenicity with this organism, although such factors as impairment of pulmonary phagocytic defense mechanisms are known to favor multiplication of P. pneumotropica (Goldstein and Green, 1967; Jakab and Dick, 1973; Jakab, 1974). This factor, at least, may be used to interpret the synergism of infections coupled with other pulmonary pathogens. An increasing body of clinical experience indicates that immunologic impairment of the rodent host predisposes to clinical expression of infection (Hajjar et al., 1991; Grossman et al., 1993; Hansen, 1995; Dickie et al., 1996; Marcotte et al., 1996; Artwohl et al., 2000; Macy et al., 2000). Thus immunocompetence of the host would seem to be as important as is pathogenicity of the organism in determining outcome of infection. Under experimental conditions, the pathogenicities of P. pneumotropica and V-factor-dependent Pasteurellaceae

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were investigated in immunodeficient F344-rnu rats (Hayashimoto et al., 2008). P. pneumotropica strains ATCC 35149 and CNP160 produced minor nasal clinical signs and histopathological changes were limited to inflammation and necrosis of the nasal mucosa; strain RPZ and the VFDP strains did not produce clinical signs or histopathological changes. Under conditions of natural infection, P. pneumotropica has been isolated most frequently among the microflora of the nasoturbinates, pharynx, conjunctiva, trachea, lungs, and uterus. Less commonly, infection of deeper organs of laboratory rats has been observed, for example, liver, spleen, and kidney (Wheater, 1967; Needham and Cooper, 1976), and P. pneumotropica has been encountered as a contaminant of MAP test tissues (Nakai et al., 2000). The earliest primary site of colonization with P. pneumotropica is the laryngopharynx (Saito et al., 1981; Mikazuki et al., 1994). Colonization of the laryngopharynx may approach an incidence of 100% in rat colonies (Saito et al., 1981), and this nidus underlies common isolation of the organism from the nasoturbinates, lower intestine, and feces. This primary localization also accords well with the analysis of common clinical presentations outlined by Besch-Williford and Wagner (1982): (1) conjunctivitis, blepharitis, and panophthalmitis (Brennan et al., 1969a,b; Wagner et al., 1969; Weisbroth et al., 1969; Rehbinder and Tschappat, 1974; Young and Hill, 1974; Kent et al., 1976; Moore and Aldred, 1978; Roberts and Gregory, 1980; Artwohl et al., 2000); (2) skin furunculosis, superficial lymph node abscesses, and mastitis (Brennan et al., 1965; Wheater, 1967; Weisbroth et al., 1969; Van der Schaaf et al., 1970; Sebesteny, 1973; Wilson, 1976; Hong and Ediger, 1978; Moore and Aldred, 1978); (3) otitis media (Wheater, 1967; Eamens, 1984; Harkness and Wagner, 1975; McGinn et al., 1992); and (4) uterovaginal infections. Besch-Williford and Wagner (1982) suggested that P. pneumotropica could ascend via the eustachian tube and nasolacrimal duct to extend infection to the middle ears and orbital structures, via oronasal contact with the genitalia to infect the preputial glands, vagina, and uterus, or by bites and suckling to infect the skin or mammaries, respectively. Reports involving inapparent infection in axenic rats indicate that the organism readily colonizes the intestines, where it may be carried asymptomatically for long periods of time (Moore et al., 1973), although enteric colonization of rats may not have epizootiologic significance beyond that of a peculiarity of gnotobiosis. In neonatal hamsters, P. pneumotropica was associated with enteritis (Lesher et al., 1985). Because of the more common association of P. pneumotropica with the respiratory system and of the difficulties of isolating P. pneumotropica from the profusion of vigorous enteric forms encountered in rat feces, it is possible that the

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epizootiological role of intestinal colonization has not been adequately investigated. In defense of a more common enteric occurrence, it has been pointed out that intranasal inoculation of fecal extracts (from infected carriers) results in respiratory infection by P. pneumotropica (Wheater, 1967). The vagina and uterus have been frequently detected as infected with P. pneumotropica (Blackmore and Casillo, 1972; Burek et al., 1972; Ackerman and Fox, 1981). Reproductive tract involvement may be either asymptomatic or clinically apparent. P. pneumotropica may be the dominant bacterial agent in the genital tract of rats (Larsen et al., 1976; Yamada et al., 1983). Studies have established that P. pneumotropica colonization of the rat uterus may approach an incidence of 60%e70% under conditions of natural infection (Graham, 1963; Casillo and Blackmore, 1972; Larsen et al., 1976). Vaginal populations of the organism have been shown to wax and wane according to stages in the estrus cycle, rising to highest levels at stages with greatest numbers of cornified, nonnucleated cells and cellular debris (Yamada et al., 1986). Transmission may take place via uterovaginal contamination (venereally), but transmission more commonly occurs by direct oronasal contact. Same cage sentinels were shown to be culture positive after 2 weeks and to have seroconverted by 3 weeks after contact exposure (Scharmann and Heller, 2001). Spread via contaminated vectors has only limited potential to spread infection. The isolate studied (NCTC8254) was quite fragile on inanimate surfaces, surviving no more than 30 min on laboratory coat fabric, about 1.5 h on plastic surfaces, and up to 2 h on mouse hair (Scharmann and Heller, 2001). Although freshly passed feces may act as a contaminated vector substance (Hong and Ediger, 1978), the practice of exposing sentinels to dirty bedding from infected carrier rodents over a period of 12 weeks did not result in infection (Scharmann and Heller, 2001). Today, most mice and to a lesser extent rats are housed in individually ventilated cage (IVC) systems to better exclude cage-to-cage pathogen contamination. Exhaust air dust (EAD)monitoring has been found to be superior to soiled bedding sentinels (SBSs) for detection of P. pneumotropica in IVC systems housing mice (Miller et al., 2016). In this study, SBSs failed to detect the organism in all cases, whereas EAD monitoring using PCR detected P. pneumotropica reliably starting at 1 week. In a similar study in which (EAD) testing at the exhaust plenum level would not be effective because of ventilated rack design, PCR monitoring of filter material placed within IVC filter tops of sentinel cages exposed to soiled bedding was found to be effective (Dubelko et al., 2018). The pathology of P. pneumotropica infection is not distinctive and resembles that caused by any of a variety

of pyogenic bacteria in similar sites. Subcutaneous abscesses of the skin, adnexal organs, or orbital structures are generally encapsulated and filled with liquid exudates. Microscopically, the lesions are suppurative with liquefactive necrosis centrally surrounded by a zone of granulomatous inflammation. Clinically silent infections in the lungs, upper respiratory tract, uterus, and intestines frequently occur without histopathologic indication of epithelial inflammation. In the lungs of mice, areas of consolidation with perivascular and peribronchial infiltration by acute and chronic inflammatory cells may be produced (Brennan et al., 1969a), although this lesion is not distinctive. Diagnosis of R. pneumotropicus and R. heylii (P. pneumotropica) can be accomplished by cultural and biochemical, serological, and PCR analyses/assays. Diagnosis of P. pneumotropica infection is established by isolation in pure culture of the organism from infected tissues. Primary isolation of the organism is accomplished readily on blood agar by using swabs from epithelial surfaces or lesion contents or aspirates from the nasoturbinates, larynx, or trachea. Isolation from feces or intestinal contents is facilitated by use of a selective medium (Garlinghouse et al., 1981; Makazuki et al., 1987). Blood agar cultures should be incubated in a moist, microaerophilic environment. After 24e48 h, colonies are 1e2 mm in diameter, nonhemolytic (although greening of the agar deep to the colonies may occur), smooth, convex, and grayish. Colonies with organisms that are Gram negative, fermentative coccobacilli, nonmotile and catalase, oxidase and nitrate positive should be regarded as presumptive for Pasteurellaceae and further identified as provisional P. pneumotropica isolates according to established biochemical profiles and properties. Many isolates, especially P. pneumotropica biotype Jawetz, fail to grow on MacConkey agar. Biotype Heyl isolates are positive for fermentation of melibiose and raffinose, whereas biotype Jawetz isolates fail to use those sugars (Nicklas et al., 1995). Mutters et al. (1995) used the reactions to lysine decarboxylase, melibiose, and arabinose to separate Heyl and Jawetzbiotypes. Definitive speciation of provisional P. pneumotropica isolates requires analysis of a panel of about 33e40 biochemical traits (Kunstyr and Hartman, 1983; Boot and Bisgaard, 1995), and biochemical reactions may be delayed and weak. Commercially available nonfermenter identification kits, for example, API20 NE and Remel RapID NF Plus, can be used to establish isolates belonging to the Pasteurellaceae, but not for speciation. Biochemical variants are common and suggestive of other, closely related Pasteurellaceae. Sensitive serologic methods such as ELISA have replaced complement fixation and agglutination tests that would not consistently detect the lower antibody levels deriving from inapparent infection (Hoag et al., 1962; Weisbroth

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et al., 1969). Moreover, the use of ELISA methodology has been demonstrated to be more sensitive than culture in the detection of clinically inapparent carriers from selected, known infected colonies of rats and mice (Wullenweber-Schmidt et al., 1988; Manning et al., 1991; Boot et al., 1995a). In this connection, both the sensitivity and specificity of detection have been improved through the use of cleaner cell wall antigens (lipooligosaccharides) than whole-cell preparations (Manning et al., 1989). The problem in recommending general use of ELISA in testing serologic unknowns derives from the issue of antigenic variability among the Pasteurellaceae generally and P. pneumotropica biotypes more specifically. There is some evidence that antigenic divergence (and some degree of host specificity) occurs among rat biotypes of P. pneumotropica, compared with those occurring in mice (Nakagawa and Saito, 1984; Boot et al., 1994a,b). Similarly, antibodies to guinea pig Pasteurellaceae, including guinea pig P. pneumotropica biotypes, do not cross-react in ELISA tests using mouse origin P. pneumotropica antigens (Boot et al., 1995b,d). The problem that results for the diagnostic laboratory is that there would appear to be no single (monovalent) serotype with antigens that would detect antibodies to all rodent biotypes, thus the potential for false negative is substantial. At the same time, the issue of common antigens among the Pasteurellaceae is widespread and underlies the potential for false positives if the objective is to serologically detect only P. pneumotropica (Wullenweber-Schmidt et al., 1988). The latter potential is sufficiently prevalent that it has been recommended that rodents older than 12 weeks are not serologically tested (Bootz et al., 1998). Identification of Pasteurellaceae at the genus and species level is now accurately accomplished by a number of molecular assays/analyses. The PCR assay developed by Bootz et al. (1998) was able to detect all 24 strains of Pasteurellaceae used in their study. The assay was 30% more sensitive than culturing, and was more specific than serologic testing. Accordingly, the primer set used in this PCR assay should be used for monitoring Pasteurellaceae infection in rodents (Boot et al., 2009). In addition to biochemical tests to differentiate R. pneumotropicus from R. heylii isolates, primer sets have been described (Kodjo et al., 1999; Wang et al., 1996). A multiplex PCR assay has been developed to allow detection of rodent Pasteurellaceae and R. pneumotropicus and R. heylii (Benga et al., 2013). MALDI-TOF analysis was able to identify, at the species level, 120 isolates representing R. pneumotropicus, R. heylii, Rodentibacter myodis, Rodentibacter rarus, Rodentibacter heidelbergensis, and Rodentibacter genomo species 2 (Adhikary et al., 2017a,b). Treatment modalities, especially for amelioration of clinical expressions of disease, have greatly improved in

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recent years. Earlier experience with antibiotics commonly resulted in clinical amelioration during treatment phases, but incomplete elimination of the organism after cessation of treatment (Spencer and Savage, 1976; Hansen, 1995). Oral dosing with enrofloxacin, a fluoroquinolone antibiotic, has been shown effective in apparently eliminating P. pneumotropica from mice without recurrence after withdrawal of the drug (Goelz et al., 1996; Ueno et al., 2002). In a later clinical study (Towne et al., 2014), enrofloxacin treatment of P. pneumotropicainfected immunodeficient NOD transgenic mice and several other infected strains was successful in eliminating the agent from the 2400 cage barrier facility. The antibiotic was given in the drinking water at a dose of 85 mg/kg body weight for a 14-day treatment cycle. At the end of 1 year, the colony remained P. pneumotropica free. The organism may be excluded from rat stocks by gnotobiotic rederivation if culturally negative uteri are used, or by embryo transfer (Reetz et al., 1988; Suzuki et al., 1996). Rodents with infected uteri may conceive and produce full-term fetuses as effectively as rats with uteri culturally free of P. pneumotropica (Flynn et al., 1968). Uterovaginal infection may result in contamination of neonates and fetuses derived by hysterectomy (Casillo and Blackmore, 1972; Hong and Ediger, 1978; Ward et al., 1978), and this potential needs to be carefully considered by those conducting rederivation programs (Reetz et al., 1988). Lack of appreciation of the clinically inapparent colonization of rat (and mouse) uterus by P. pneumotropica more than any other single factor has been responsible for the reintroduction of this organism to isolators or barriers of gnotobiotic rodents, and has been a frequent cause of barrier “breaks” (Blackmore, 1972; Casillo and Blackmore, 1972). Enrofloxacin has been used preparatory to rederivation to clear or reduce uterine populations of P. pneumotropica to enhance the likelihood of a successful outcome (Macy et al., 2000). 2. Muribacter muris (Actinobacillus muris) Earlier studies had shown that A. muris was a species unrelated to other members of Actinobacillus and that A. muris was unrelated to P. pneumotropica (Piechulla et al., 1985; Ryll et al., 1991). A study to further investigate the taxonomy of A. muris was done on 130 strains for genotypic characterization by 16S rRNA and partial rpoB gene sequencing, with whole-genome sequencing done on the type strain. As determined from this study, a new genus and species, M. muris, were proposed based on a distinct phylogenetic position with major divergence from the existing genera of the family Pasteurellaceae (Nicklas et al., 2015). Accordingly, A. muris (Bisgaard, 1986) has been reclassified as M. muris.

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Increasingly, the rodent actinobacilli are being diagnostically encountered, usually in asymptomatic carriers, but occasionally in association with clinical conditions of the respiratory and reproductive systems. The 2014 FELASA report (Mahler et al., 2014) recommends that Pasteurellaceae other than P. pneumotropica should be screened according to the specific needs of an institution. Several Actinobacillus species have been reported from laboratory rodents, often in coinfection with P. pneumotropica. In a survey of feral rodents, many were found to be reservoir hosts for Actinobacillus species, P. pneumotropica, or both (Boot et al., 1986). The earliest reference to rodent actinobacilli is that of Simpson and Simmons (1980), who reported isolation of two species, Actinobacillus equuli and one not further identified, from nasopharyngeal washes and ceca of laboratory rats and hamsters undergoing health surveillance screening tests. One of the hamster isolates originated from a cheek pouch abscess, but the rats and other hamsters were asymptomatic. In the same year, Lentsch and Wagner (1980) reported isolation of A. equuli and Actinobacillus lignierseii from oropharynx, conjunctiva, and middle ears of laboratory rats, mice, and guinea pigs. Shortly after, Pasteurella ureae was reported as a copathogen with P. pneumotropica in an epizootic of mouse reproductive tract disease (Ackerman and Fox, 1981). The Ackerman and Fox isolate was later reclassified by Bisgaard (1986) as A. muris in the original description of this new species. The species name Actinobacillus ureae is retained to identify a different organism whose natural distribution appears limited to humans (Mutters et al., 1984). Also in 1981, an organism later reclassified as A. muris was isolated from instances of mastitis in laboratory mice (Gialamas, 1981). The taxonomic position of Haemophilus influenzaemurium is unsettled. The name H. influenzae-murium is no longer listed as a valid name (Euzby, 2003). Based on DNA homology, classified this organism with A. muris, and for that reason it is included in this section. The closest related taxon outside the M. muris group based on rpoB sequence comparison was H. influenzaemurium, which is at the lower range of similarity between species of the Pasteurellaceae. The 16SrRNA gene sequence similarity was also below the species level. Accordingly, further taxonomic investigation will need to be done to show whether H. influenzae-murium belongs to Muribacter as a genomospecies (Nicklas et al., 2015). H. influenzae-murium was first isolated from mice with respiratory disease (Kairies and Schwartzer, 1956; Ivanovics and Ivanovics, 1937) and was not reported again for 40 years until encountered during health surveillance tests on healthy laboratory

mice in Hungary, reisolated, and characterized (Csukas, 1976). The primers used by Bootz et al. (1997, 1998) to detect all members of the Pasteurellaceae detect all reference strains of rodent actinobacilli. The primers used by Wang et al. (1996) and designed to detect P. pneumotropica would not detect A. muris if a more stringent annealing temperature (57
C) were used (Weigler et al., 1998). 3. Rodentibacter heidelbergensis and Rodentibacter trehalosifermentans (Haemophilus) Recently, the classification of Haemophilus (V-factor dependent) has been changed to two species: R. heidelbergensis and R. trehalosifermentans (Adhikary et al., 2017a,b;Boot et al., 2018). Both the incidence and importance of the two Rodentibacter spp. (Haemophilus) as pathogens of rats are difficult to assess. They appear to be only weakly pathogenic and are usually clinically inapparent; however, the 2014 FELASA report (Mahler et al., 2014) indicates they should be screened according to the surveillance needs of an institution. The Haemophilus species are fastidious in their growth requirements, and microbiologic screening for them requires an additional sequence of media. Their isolation and identification pose a significant challenge for clinical microbiologists. The incidence of Rodentibacter in laboratory rats may be substantially underestimated because the usual diagnostic regimen does not use the media required for their isolation. One survey (Nicklas and Benner, 1994) indicated an incidence of 21% in rat diagnostic specimens. The V-factor-dependent Pasteurellaceae have been encountered in episodes of respiratory disease in rats often under circumstances where a diagnostic search for X-factor- or V-factor-dependent pathogens was prompted by failure to isolate an etiologic agent on blood agar (Nicklas, 1989) or by failure to isolate Pasteurella in rat sentinels that had seroconverted to positive against P. pneumotropica antigens (Boot et al., 1995c). Moreover, intercurrent infection by Haemophilus can modulate certain research protocols. It has been demonstrated that infection has a depressive effect on development of adjuvant arthritis in LEW rats (Boot et al., 1995a). Clinical specimens for isolation are essentially the same as those recommended for P. pneumotropica, that is, nasopharyngeal washes or swabs of the oropharynx or vagina. The rodent Haemophilus species are V-factor dependent for their growth and will not be isolated from clinical inocula cultured on (unsupplemented) blood agar. The required V- (NAD) and X- (hemin)

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factors are both provided by the gentle heating of blood cells in the preparation of chocolate agar, the medium recommended for clinical isolation. Supplementation of the medium with clindamycin can be used to inhibit overgrowing microflora (Garlinghouse et al., 1981; Boot et al., 1995c). Inoculated chocolate agar plates should be incubated in a moist, microaerophilic (5% e10% CO2) environment for 24e48 h. Dependence on V- or X-factors, or both, for growth can be determined by the growth patterns of lawns of the isolate streaked onto MuellereHinton agar plates, onto which discs with V- or X-factors or both (VeX) are placed. Radial growth, for example, surrounding the V and XeV discs are indicative of V-factor dependence. Caution should be exercised in assigning V-factor-dependent Pasteurellaceae to the genus Haemophilus, however, because there are X-factor- and V-factor-dependent strains of both Pasteurella and Actinobacillus (Nicklas et al., 1993; Nicklas and Benner, 1994; Boot et al., 1995a). The transmissibility of V-factor-dependent Pasteurellaceae between laboratory rodents, including rats, mice, and guinea pigs, has been demonstrated (Boot et al., 2000); thus the separation of host species having disparate pathogen profiles is recommended. Presumptive Haemophilus isolates may be further characterized by use of commercial kits, for example, API NH (bioMerieux France), with the proviso that it be recognized that profiles or software supplied by the kit are oriented to identification of human isolates and are generally inadequate for rodent isolates (Brands and Mannheim, 1996). Rat Haemophilus isolates will usually be classified by the API NH system as H. parainfluenzae (Boot et al., 1995a, 1997). Biochemical profiles for rodent Haemophilus species are tabulated in the following references: Brands and Mannheim (1996), Boot et al. (1997, 1999), and Hansen (2000). Rodent Haemophilus isolates may be detected in swab samples from infected rats by PCR; however, the primers reported by Bootz et al. (1997, 1998) detect all members of the Pasteurellaceae and thus cultural criteria would be needed to confirm PCR positives. PCR negatives would be acceptable to demonstrate that the samples were free of Pasteurellaceae, including Haemophilus sp. It has been found that rat strains differ in antibody response to natural Haemophilus spp. infection (Boot et al., 2005). Because of a one-way cross-reaction, antibodies to many Haemophilus strains sufficiently react with P. pneumotropica ELISA antigen to enable detection of Haemophilus infection, although the converse is not necessarily true (Boot et al., 1995c). As mentioned earlier, this predilection also limits specificity of serologic screening tests to detect P. pneumotropica. In their study of Haemophilus strains, Boot et al. (1997) demonstrated that a panel of at least two antigenic types

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(P. pneumotropica and the H21 strain of Haemophilus) would be needed to reliably detect all strains of Haemophilus. Thus PCR may be a simpler and more costeffective way of screening rat populations for presumptive Haemophilus status. More recently, MALDI-TOF has been reported to be capable of identifying R. heidelbergensis at the species level (Adhikary et al., 2017a,b).

K. Opportunistic Bacterial Pathogens A number of bacterial genera can be considered as opportunistic (potential) pathogens in laboratory rats as well as for other animal hosts, including humans. Factors that affect their bacterialehost interactions and subsequent degree of pathogenicity in rats include the host immunestatus, the microbiome diversity and relative dominance of a specific microorganism within a tissue/organ system of the host, age of the host, environmental or experimentally related stressors, and virulence factors specific to the agent infecting the host. The exclusion of specific bacterial species that may be potential pathogens and/or undesirable study variables will depend upon the specific requirements of investigators and institutional resources. To assist institutions in designing effective infectious agent monitoring programs, FELASA periodically publishes recommendations for health monitoring of rodent and rabbit colonies (Mahler et al., 2014, and for a chronologic summary, see Table 12.1). For bacterial agents that occur very infrequently or that can be designated as opportunistic pathogens, FELASA recommends that testing and the frequency of testing should be done predicated on institutional/investigator needs. The bacterial species listed by FELASA in this category include Bordetella bronchiseptica, C. kutscheri, Klebsiella oxytoca, Klebsiella pneumoniae, P. aeruginosa, and Staphylococcus aureus; FELASA notes that this list is not inclusive. A recurring question of significance when an opportunistic bacterial species has been isolated is whether one has isolated an agent inducing a potentially pathogenic infection or only an asymptomatic colonization. For health surveillance purposes, all of this group should be considered reportable. The possible introduction of these agents in laboratory rats reinforces the principle of separation of species and rats of disparate microbial profiles within laboratory animal resources. 1. Klebsiella pneumoniae and Klebsiella oxytoca A large-scale health surveillance report indicated a very low prevalence for both K. pneumoniae and K. oxytoca in laboratory rats (Pritchett-Corning et al., 2009). The data were derived from bacterial testing done by a major commercial rodent diagnostic laboratory for clients that were pharmaceutical and

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biotechnology companies, universities, and government laboratories. In their report, data derived from culturetesting over 3-year (Europe) and 5-year (North America) periods reflected a prevalence of 0.0% and 0.55%, respectively, for K. pneumoniae (N ¼ 7513) and 0.0% and 0.37% for K. oxytoca (N ¼ 7315). K. pneumoniae can be isolated from the rat intestinal tract in the absence of clinical signs or pathology. There are very few reports of naturally occurring clinical diseases due to the opportunistic pathogens K. pneumoniae and K. oxytoca (Hansen, 1995). An epizootic was reported in which K. pneumoniae was responsible for clinical disease in 82 of 2300 Wistar rats of an institutional-based breeding colony (Jackson et al., 1980). These affected rats displayed unilateral and bilateral soft subcutaneous masses of the submandibular, parotid, and inguinal areas. The skin overlying these lesions, in some cases, had ruptured, discharging purulent exudates. The foregoing masses represented abscessed lymph nodes and associated connective tissue. The infections in some rats were disseminated with splenic, hepatic, renal, lung, and mesenteric nodes containing lesions. K. pneumoniae serotype 5 was isolated from the involved lesions and abscesses. An untypable K. pneumoniae was isolated from feces of affected and nonaffected rats, but not from lesions in any animals. One report cites K. pneumoniae in nude rats (LEW/Mol-rnu/rnu)(Hansen, 1995). This nude rat colony had been medicated with multiple antibiotics for 30 weeks primarily to suppress the primary target P. pneumotropica. This treatment was successful only until therapy ended. Although K. pneumoniae had not been previously isolated from rats in the colony, it emerged as a prominent intestinal isolate postantibiotic treatment. Clinical signs or lesions were not reported with the infection. Despite the ubiquitous distribution and the capabilities of Klebsiella spp. as opportunists, well-designed microbial surveillance programs and effective husbandry protocols have minimized colonization of Klebsiella in immunologically impaired rats and mice. Klebsiella species are Gram-negative, short rod, nonmotile, capsulated, facultatively anaerobic members of the Enterobacteriaceae. They are readily isolated from enteric sequence cultures made from feces or oropharyngeal swabs and less frequently from nasopharyngeal washes. Primary isolation media include Gram-negative broth, MacConkey agar, or blood agar. A selective medium has been developed to facilitate clinical isolation of this organism (Kregten et al., 1984). The IMViC reactions may be used to distinguish K. pneumoniaee e þ þ) from K. oxytoca (þe þ þ) although, in practice, clinical microbiologists simply assign indole positive rodent Klebsiella isolates to K. oxytoca. Commercial identification kits, for example, API 20E, are widely used to identify presumptive Klebsiella isolates to species.

The polysaccharide capsular material of Klebsiella can be typed into one of six types and is known to be an important modulator of virulence (Held et al., 1992), but it appears to be of limited value as an epizootiologic tool (Davis et al., 1987). Both rats and mice are used extensively as research models for Klebsiella infection (for summaries, see NRC, 1991; Baker, 1998). 2. Bordetella bronchiseptica Today, there is negligible evidence that B. bronchiseptica is responsible for naturally occurring disease in laboratory rats. Moreover, in a wide-scale study on the prevalence of infectious agents in rats, culturing results from over 8200 rats showed a 0.0 prevalence of B. bronchiseptica (Pritchett-Corning et al., 2009). It is important to note, however, that the majority of samples tested by a large commercial diagnostic laboratory were from pharmaceutical and biotech companies, not from academic and governmental sources in North America and Europe. Marked decreases in the reported cases of B. bronchiseptica over the past 50 years have led to changes in recommendations for the frequency of monitoring for this bacterial agent (Table 12.1). The 2014 FELASA working group on health monitoring recommendations for B. bronchiseptica places the agent in a group for which monitoring is optional, and if pursued the frequency would depend upon local circumstances (Mahler et al., 2014). This is a marked change from the FELASA working group’s 2002 report that recommended a testing frequency of 3month. Moreover, the National Research Council placed B. bronchiseptica among agents that are not conclusively demonstrated to be natural pathogens of rats and mice (NRC, 1991). Bemis et al. (2003) presented an excellent overview of reports that dealt with B. bronchiseptica infection in rats and mice and noted that there have been few reports of naturally acquired disease due to this microorganism since 1960. It is very likely that in many of the early reports, coinfection with other primary pathogens such as M. pulmonis and Sendai virus contributed greatly to the observed disease and pathology. Winsser (1960) described lesions in rats from which B. bronchiseptica was recovered; however, the rats in those studies were conventional, and the role assumed for B. bronchiseptica cannot with confidence be attributed solely to that organism. In recognition of the ambiguity of the Winsser study, Burek et al. (1972) explored the experimental pathogenicity of B. bronchiseptica for laboratory rats. The organism induced acute to subchronic bronchopneumonia in both germ-free and conventional rats. The pathology of experimental infections is described (Burek et al., 1972; Percy and Barthold, 2001). In contemporary well-managed rodent resources there is very little likelihood that interspecies transfer of the organism would occur. However, B. bronchiseptica is a frank pathogen of

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guinea pigs and this species could serve as a potential infectious reservoir to other rodent species. 3. Staphylococcus aureus S. aureus is a commonly occurring commensal in essentially all animal species, including humans. Staphylococcus may be isolated from the oropharynx, intestinal tract, and skin of asymptomatic carrier rats. Although it is very infrequently associated with clinical disease in immunocompetent rats, S. aureus has long been associated with ulcerative dermatitis of laboratory rats (Ash, 1971; Fox et al., 1977; Wagner et al., 1977; Galler et al., 1979; Kunstyr et al., 1995) in which inciting factors such as immunodeficiency, protein nutritional deficiency, and trauma due to fighting or arthropods are involved. This ulcerative dermatitis occurs episodically, is not age or stock/strain dependent, and usually does not have an incidence of more than 1%e2% of the animals within a colony. Skin lesions are most often located on the shoulders and lateral cervical regions, and typically present as areas of alopecia with irregular, reddish, weeping, or crusty ulcerating skin. S. aureus is consistently isolated from such lesions, and characteristic microcolonies of staphylococci are observed microscopically in the superficial debris. Illustrations of the gross and microscopic appearance of these lesions may be found in reports by Fox et al. (1977) and Wagner et al. (1977), and texts by Percy and Barthold (2007) and Barthold et al. (2016). In a relatively recent report on the prevalence of infectious agents in laboratory mice and rats (PritchettCorning et al., 2009), S. aureus was isolated from 23.6% of 7365 rats that were screened. These animals/specimens were primarily from pharmaceutical and biotech companies in both North America and Europe with the remainder from academia and governmental institutions. The most recent FELASA recommendations for health monitoring of rodents and rabbits placed S. aureus in an agent group for which monitoring is optional and dependent on specific institutional or investigative needs. In a later report (Marx et al., 2017) based on data from 63 US institutions there were only two instances of Staphylococcus outbreaks over a 2-year period. Of these 63 institutions only 7% excluded Staphylococcus from their rat and mouse colonies. Rats that are immunodeficient should be monitored and maintained S. aureus-free by enforcing bioexclusion husbandry practices to prevent invasive and systemic infections (BeschWilliford and Wagner,1982). A heritable component to ulcerative dermatitis seen in Swiss Webster mice and certain inbred strains (Clarke et al., 1978; Shults et al., 1973; Wardrip et al., 1994; Slattum et al.,1998) has not been reported as a factor in epizootic occurrences in rats. The organism is easily cultured on blood agar media from the nasopharynx, skin, or lesions. Isolates of

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S. aureus are Gram-positive cocci and occur as grapelike clusters, are coagulase and catalase positive, and produce yellow pigmented colonies with a-or b-hemolysis on blood agar media. Culturing on mannitol salt agar is a useful selective medium and differentiates S. aureus from S. epidermidis. Further identification of isolates may be done by phage typing, commercial biotype kits, or 16S rDNA typing. Staphylococci are quite resistant to environmental temperature extremes and to drying, and are easily vectored by contaminated clothing, ungloved hands, and respiratory droplets (Blackmore and Francis, 1970). Since very few institutions exclude S. aureus from their mouse and rat colonies (Marx et al., 2017) it is not surprising that the organism is frequently present asymptomatically from the intestinal tract, skin, or nasopharynx in the immunocompetent rat. About 30% of humans carry S. aureus in their nasopharynx and on their skin and thus human-to-rat transmission is likely a significant factor for the introduction and prevalence of the organism in rat colonies. Greater emphasis on biosecurity measures such as personnel donning disposable gloves, gowns, and face masks when handling Staphylococcus-naive animals and use of microisolator cages and cage-changing stations is a requirement to help limit transfer from personnel and fomites to rat colonies for institutions choosing to exclude the organism.

III. MYCOPLASMAL INFECTIONS A. Introduction The genus Mycoplasma is one of eight genera within the Class Mollicutes. Mycoplasma are bacteria that lack a cell wall and, with a 0.2-mm diameter, they represent the smallest free-living prokaryotes known. Their genome may be as small as 600 kb, one-tenth that of Escherichia coli. Accordingly, Mollicutes are often models for studying the minimal metabolism necessary to sustain independent life. They lack cytochromes or the tricarboxylic acid cycle except for malate dehydrogenase activity and are unable to synthesize pyrimidines or purines de novo (Pollack et al., 1997). The first Mycoplasma species identified was the causative agent of contagious bovine pleuropneumonia and was successfully cultivated on artificial media by Nocard and Roux in 1898. Later, microbial agents with similar characteristics were isolated from a wide range of animal species. These isolates were named PPLOs by Klieneberger (Klieneberger, 1938; Klieneberger and Steabben, 1940), and this term was commonly used in the literature until the mid-1960s. Mycoplasma are known to be significant pathogens in most species of animals as well as in humans. Generally,

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they have tropisms for the respiratory tract, urogenital tract, or articular cartilage/synovial tissue in the animal or human host. Their pathogenicity is typically associated with a chronic disease course rather than a fulminant course. In laboratory rats, three Mycoplasma species, M. pulmonis, Mycoplasma arthritidis, and Mycoplasma haemomuris (formerly classified as Haemobartonella muris), have been shown to be pathogenic; however, M. pulmonis is the most significant pathogen of the three. Until 1969, the term chronic respiratory diseasewas used to describe a respiratory disease in rats believed at the time to be due to two causative coinfective agents: a virus that produced bronchiectasis and M. pulmonis that induced an upper respiratory tract catarrhal infection (Nelson, 1951). Kohn and Kirk (1969) first demonstrated that M. pulmonis was both necessary and sufficient to account for all features of the upper and lower respiratory tract disease syndrome. In the next several years, other reports confirmed this finding and significantly expanded information on the pathogenesis of this Mycoplasma (Kohn, 1971a,b; Lindsey et al., 1971; Jersey et al., 1973; Cassell et al., 1973).

B. Mycoplasma pulmonis (Respiratory Mycoplasmosis) Murine respiratory mycoplasmosis was arguably one of the most significant infectious diseases in laboratory rats until the 1980s, at which time commercial sources of rats began offering stock free of M. pulmonis. Today, M. pulmonis infections are essentially limited to rats produced by noncommercial institutions, rats that become infected through contact with such breeding colonies, or rats that have been inoculated with contaminated biological materials. The very low prevalence of M. pulmonis in laboratory rats today is reflected in that of over 81,000 serological samples received by a commercial rodent diagnostic laboratory (Pritchett-Corning et al., 2009), only 0.16% of the samples were positive from North American organizations and 2.57% from those in Europe. By PCR, the prevalence in samples from North America was 0.29%. However, it should be noted that most of the samples tested were from pharmaceutical and biotechnology organizations with much smaller numbers from academia and governmental institutions. Clinical signs in rats infected with M. pulmonis are quite variable; in many cases, no signs are present even though significant pulmonary lesions may exist. The prevalence and severity of signs typically increase with the age of the rat and the presence of experimental or environmental stresses placed upon the animal. Clinical signs include sniffling, rales, roughened haircoat, dyspnea, weight loss, and red staining of the eyes periorbitally and at the nares caused by increased porphyrin

secretion of the Harderian glands. Not uncommon is torticollis associated with extension of the infection from the nasal mucosa through the eustachian tube to the middle and inner ear. Mucopurulent exudation occurs in the nasal cavity and trachea; however, it may not be readily evident. The pulmonary lesions vary from small surface areas that are dark red or gray to entire lobes that are affected. The apical lobe and cranial portion of the left lobe tend to be more often affected than are the other lobes; however, distribution varies among animals. The typical end-stage lung lesion is a cobblestone-like surface, reflecting marked bronchiectasis. The size of these abscessed bronchi varies from several millimeters to coalesced abscesses up to 1 cm in diameter. The lung surface surrounding these multiple yellowish lesions is dark because of consolidation and atelectasis. These typical gross lesions are well illustrated in the text by Barthold et al. (2016). Cross-sections of affected lungs reveal markedly dilated bronchi impacted with caseated exudates and consolidation of varying amounts of parenchyma. The inflammatory response to M. pulmonis infection is variable but can be intense, particularly after an extended period of time. The response at all sites in the respiratory tract and middle ear is characterized by a submucosal lymphocytic infiltrate and primarily a neutrophilic leukocyte exudate at the epithelial cell surface. Within the lung, peribronchial lymphoid hyperplasia is the initial dominant lesion, and as it increases, leukocyte infiltration of the bronchial and bronchiolar lumens leads to bronchiectasis and bronchiolectasis. Mucosal epithelial cells often become cuboidal or squamoid in bronchiectatic airways, and hyperplasia of the epithelial mucosa may occur. Another outcome of M. pulmonis infection is angiogenesis and blood vessel remodeling associated with microvascular leakiness due, in part, to formation of endothelial gaps. This is accompanied by an influx of inflammatory cells, epithelial thickening, mucous gland hypertrophy, fibrosis, and an abnormal sensitivity to substance P (Kwan et al., 2001). M. pulmonis is transmitted vertically and horizontally within rat colonies. It is readily transmitted from infected to noninfected rats by direct and indirect contact. However, the rapidity of transmission may vary within colonies because of differences in room ventilation rates, husbandry practices, and animal density. The importance of fomites in transmission is not well defined. The organism does not survive long periods outside the host; however, personnel and contaminated equipment may be of importance in some instances. Within breeding colonies, dams may infect their litters postpartum or in utero because the female genital tract is frequently a site of infection. Most significant in the

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epizootiology of M. pulmonis infection is that infected rats typically remain as a life-long source for transmission to naive rats of any age. The most important factors that modulate the pathogenesis M. pulmonis are (1) those effecting cytadherence, (2) genotype of the rat, (3) coinfection with a virus pathogen, and (4) environmental influences. A summary of studies that have dealt with elucidation of these modulating factors is given next. Mycoplasma spp., in general, colonize the epithelial cell surfaces of the respiratory and urogenital tracts. Unlike most other bacteria, Mycoplasma lack fimbriae that are involved in adhesion to the host cell membrane. Cytadherence of Mycoplasma is associated with adhesins located within the cell membrane that attach to specific receptors on the host cell membrane (Razin and Jacobs, 1992). M. pulmonis has a tropism for the ciliated epithelial cells of the respiratory tract and the genital tract in rats. Electron micrograph illustrations showing this tropism may be seen in the report of Kohn (1971a) and in the text by Barthold et al. (2016). Under experimental conditions it also cytadsorbs to the ciliated epithelial surface of ependymal cells (Kohn and Chinookoswong, 1981). Absence of a cell wall imparts a unique cell membrane-to-cell membrane relationship, allowing for transfer of needed hostmetabolites to the microorganism and exposure of the host cell to Mycoplasma-generated cytopathological metabolites such as peroxide and superoxide radicals (Razin and Jacobs, 1992). Some Mycoplasma, most notably the human pathogen M. pneumoniae, have an attachment organelle that aligns the organism with the host cell receptor site (Collier and Clyde, 1971). Adherence of M. pulmonis to the host cell membrane is key to the pathogenesis of respiratory disease, and the degree to which the organism successfully adheres to the nasal mucosa is likely of major significance to the outcome of infection in the lower respiratory tract (Schoeb et al., 1993a). Using tracheal organ culture, Schoeb et al. (1993b) studied the modulating effects that sialodacryoadenitis virus (SDAV) and Sendai virus infections, age of the host, vitamin A deficiency, and exposure to ammonia may have on adherence caused by alteration of the respiratory mucosal surface. In their in vitro model, these investigators found that prior exposure of rats to SDAV and to vitamin A deficiency substantially increased the number of organisms adhering to the tracheal explants derived from the animals. It is known that mouse genotype influences respiratory disease susceptibility to M. pulmonis (Lai et al., 1993; Cartner et al., 1996). In rats, marked susceptibility differences exist between LEW and F344 strains. After experimental infection with M. pulmonis, LEW rats mount a significantly increased inflammatory response to the

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organism in comparison to F344 rats (Davis et al., 1982; Davis and Cassell, 1982). Lung lesions in LEW rats developed earlier after infection, became more severe, and were characterized by larger areas of bronchusassociated lymphoid tissue hyperplasia. In both Lewis and F344 rats, the upper respiratory nodes were determined to be the initial site of antibody production after experimental infection (Simecka et al., 1991), and a major site of antibody-forming cells through a 28-day study. By 14 days after infection, antibody-forming cells were found in other tissues, suggesting that the upper respiratory tract may be a significant source of primed or activated B cells that migrate to the infected tissues. In the same study, there was a major difference in the number of M. pulmonis-specific B-cells in the draining lymph nodes of the lungs, with LEW rats having a larger proportion. There was also a greater antibody response in the lungs and upper respiratory nodes of LEW rats. The investigators noted that this may indicate that antibody production in the upper and lower respiratory nodes contributes significantly to the differences in response to infection between the two strains. Serum antibody levels alone do not resolve lesions and are not responsible for differences in lesion severity between the two rat strains (Simecka et al., 1989). The pathogenicity of M. pulmonis is modulated by coinfection with viruses. Schoeb et al. (1985) found that gnotobiotic F344/N rats experimentally infected with Sendai virus and M. pulmonis had more severe lesions and a greater proliferation of M. pulmonis than did rats infected with only M. pulmonis. Tracheal mucosa in the coinfected rats was nearly destroyed by severe lymphocyte and plasmacyte accumulations, whereas monoinfections induced mild inflammatory changes. At 3e4 weeks after infection, lungs from coinfected rats had suppurative pneumonia, atelectasis, and, in some cases, bronchiectasis, whereas lungs from mono-infected rats had essentially normal lungs. Similarly, Schoeb and Lindsey (1987) observed that F344/N rats coinfected with SDAV developed more severe lesions throughout the respiratory tract than animals monoinfected. Although the mechanism for this enhanced pathogenicity is not known, the investigators suggested that coinfection with a virus such as Sendai virus or SDAV could induce immunosuppression, inhibition of macrophage function, or increased cytadhesion of M. pulmonis. However, it has been shown that coinfection with either Sendai virus or SDAV does not alter intrapulmonary killing or physical clearance of M. pulmonis in either LEW or F344 rats (Nichols et al., 1992). Broderson et al. (1976) first reported the direct association of increased intracage ammonia levels on severity of rhinitis, otitis media, tracheitis, and pneumonia in rats infected with M. pulmonis.Pinson et al. (1986) examined

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the nasal mucosa of rats infected with M. pulmonis exposed to an ammonia level of 100 ppm. Within 9 days of ammonia exposure, more severe degenerative changes occurred in the mucosa infected with Mycoplasma than in the mucosa of noninfected rats. Exposure to ammonia in the absence of Mycoplasma infection induced only mild inflammatory changes, whereas widely distributed hyperplastic, metaplastic, and inflammatory changes were induced in Mycoplasmainfected rats. In another report (Schoeb et al., 1982), F344 rats were exposed to 100 ppm of ammonia and inoculated intranasally with M. pulmonis. This resulted in an increase in M. pulmonis titers and an exacerbation of lung lesions. In addition, it was determined that ammonia is almost entirely absorbed in the nasal passages, not in the trachea or lungs. This finding indicated that the increased numbers of Mycoplasma in the trachea and lung are probably a secondary effect rather than a direct effect of ammonia levels at either site. ELISA testing is the most frequently used method to determine if the organism is present within a rat colony. A presumptive diagnosis is difficult to make based only on characteristic gross lung lesions associated with M. pulmonis because other pathogens such as the CAR bacillus and C. kutscheri induce similar lesions. Culturing the organism from sites such as the nasopharynx, middle ear, trachea, lung, and oviduct is a reliable means to define the M. pulmonis status of an animal or a colony (Cassell et al., 1983). However, the nutrient requirements for Mycoplasma are more complex than for most other bacteria because Mycoplasma require a source of sterol, usually supplied by addition of either sterile horse or swine serum to the broth and agar media. Accordingly, ELISA testing has replaced culturing as the principal means to diagnose infections because it is less time consuming and costly and is comparable in reliability (Mia et al., 1981; Cassell et al., 1986, Lindsey et al., 1991). PCR is a sensitive means to diagnose M. pulmonis infections that may be below the threshold for detection by culture and ELISA testing, to confirm serological results and to screen immunodeficient strains/stocks. PCR detects M. pulmonis in either nonfixed or paraffinembedded tissues (Van Kuppeveld et al., 1993; Brunnert et al., 1994; Sanchez et al., 1994; Schoeb et al., 1997b). An excellent review of laboratory methods to diagnose mycoplasmosis in laboratory rodents is presented by Davidson et al. (1994). Using mice, Longanbill et al. (2005) developed an M. pulmonis fluorogenic nuclease assay that detected the equivalent of 20 viable organisms. Although M. pulmonis very infrequently occurs in laboratory rat colonies today, regular testing for this pathogen must be part of any screening program because the organism continues to exist in a small subset of

conventional colonies of rats and mice, in wild or feral rodents, in contaminated biological materials, and is widely prevalent in pet rodents. Once M. pulmonis is introduced into a colony, there is no effective means to eliminate it other than through rederivation by cesarean section or embryo transfer and removal of all animals shown to be infected. If cesarean section is the method chosen, the uterus and oviducts of the donor dam must be tested for M. pulmonis by culturing and/or PCR and the rederived litter maintained in quarantine until determined to be M. pulmonisfree by repeated ELISA and PCR testing. Antibiotic therapy is not effective in eliminating the organism from infected animals and immunization protocols have been investigated but were found to be nonprotective (Cassell and Davis, 1978). Before the elimination of M. pulmonis from commercial sources of laboratory rats, this organism was responsible for catastrophic effects on many long-term toxicologic, gerontologic, and cancer studies. Today, M. pulmonis remains in some conventional rat and mouse colonies; these sources and contamination of M. pulmonis-free colonies by feral or wild rodents still pose risks to long-term studies if appropriate measures are not taken to exclude the organism from barrier facilities. M. pulmonis infection is incompatible with essentially any investigation in which the respiratory or female genital tract is involved. A number of studies have shown its effect on the immune system. These include suppression of humoral antibody response to sheep erythrocytes, modulation of collagen-induced arthritis, and induction of nonspecific mitogen effects on lymphocytes (Cassell et al., 1986; Lindsey et al., 1991).

C. Mycoplasma pulmonis (Genital Mycoplasmosis) Although M. pulmonis is most often associated with respiratory infections, the organism has a strong tropism for the epithelia of the female genital tract. In female rats that have naturally occurring respiratory mycoplasmosis, up to 40% will also have M. pulmonis-infected genital tracts. Of these, it is estimated that about 30% will have grossly evident oophoritis and salpingitis (Cassell et al., 1979). Infection of the female genital tract is primarily a function of colonization of the vagina with subsequent ascending spread to the uterus and oviducts. However, it is possible that hematogenous spread to the female genital tract may also occur in naturally occurring infections since intravenous inoculation of M. pulmonis results in a very high rate of oviductal and uterine infection. Infection of the genital tract in male rats has been reported. M. pulmonis has been isolated from the testes, epididymides, and vas deferens of rats that were bred to genital tract-infected females. Infection of

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the male genital tract may be a factor associated with infertility (Steiner and Brown, 1993), and it has also been shown in vitro that the organism adheres to spermatozoa (Cassell et al., 1981b). Clinical signs associated with genital mycoplasmosis are limited to infertility and to fetal and neonatal deaths. The most frequently occurring lesions of the female genital tract are purulent exudates within enlarged oviducts and distended ovarian bursas caused by exudation. Uteri may not show any gross abnormalities; however, in some cases pyometra occurs (Graham, 1963; Cassell et al., 1979). The inflammatory response in the oviduct is characterized by a polymorphonuclear cell exudate within the lumen, hyperplasia, and metaplasia of the ciliated columnar epithelium and a submucosal lymphoid infiltrate. This response quite closely mirrors that seen with M. pulmonis-infected trachea and bronchi. Within the uterus, a polymorphonuclear cell inflammatory response occurs with varying intensity. Similar to infection in the respiratory tract, the organism adheres to the cell surface without migration into the submucosa. In the gravid uterus, there are foci of polymorphonuclear cells in the decidual and basal layers of the placenta (Steiner and Brown, 1993). M. pulmonis induces a placentitis similar to that described in deciduitis of humans and represents a model system for investigations into the pathogenicity of uterine infection (Peltier et al., 2003). A number of studies have been done to determine factors involved in outcomes associated with genital tract infections (Brown and Steiner, 1996). The pathogenesis associated with fetal death has been studied by experimentally inoculating rats intravaginally with M. pulmonis before breeding. Subsequent to inoculation, the organism infects the uterus, invading the placenta and establishing an infection in the amnion cavity at day-14 gestation. At gestation day 18, it can be isolated from the oropharynx and lungs of fetuses (Steiner et al., 1993). Similarly to respiratory mycoplasmosis, rat stocks/strains differ in their susceptibility to genital tract infection and pregnancy outcome. A study comparing infection outcome of Sprague-Dawley, Wistar, F344, and LEW rats found that Sprague-Dawley rats had a greater prevalence of infertility, decreased litter size, decreased pup birth weight, increased resorptions and stillbirths, and the highest rate of pulmonary infection in neonatal pups. LEW dams, at 1-day postpartum, had the lowest genital tract infection rate, and their litters lacked evidence of vertical transmission of M. pulmonis. In this study, intravaginal inoculation of the organism resulted in self-limiting genital tract infections in the LEW and F344 strains (Reyes et al., 2000). The hormonal influence of testosterone treatment on increasing the prevalence and severity of M. pulmonis-

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induced genital mycoplasmosis has been reported (Leader et al., 1970). Cesarean section is commonly used to rederive a rat line to render it free of M. pulmonis. If the genital tract or the fetuses of the rat undergoing cesarean section were infected with the organism, it is obvious that the success of the process to rederive a litter free of Mycoplasma would most likely be compromised. Graham (1963) was perhaps the first to caution that vertical transfer of M. pulmonis via infected uteri of hysterectomized rats could lead to unsuccessful rederivation of litters. Vertical transmission of the organism has been reported in germ-free rats that were contaminated by hysterectomized dams that were infected with M. pulmonis (Ganaway et al., 1973). Accordingly, it is essential to use ELISA and PCR testing to ensure that rederived litters are, in fact, free of Mycoplasma.Rederivation by early embryo transfer is an alternative to cesarean section. Trypsin treatment of embryos has been successfully used to help ensure that Mycoplasma and viruses are not contaminants of transferred embryos (Rouleau et al., 1993).

D. Mycoplasma arthritidis (Articular Mycoplasmosis) Klieneberger (1938) isolated a PPLO that she termed L4 from the middle ear and oropharynx of laboratory rats. The L4 organism was subsequently reclassified as Mycoplasma arthritidis. Although rarely a cause of naturally occurring arthritis in laboratory rats, the organism has been reported in earlier surveys to infect conventionally maintained colonies and has been reported in barrier-maintained rats (Cox et al., 1988). Today, all commercial sources of barrier-maintained rats in the United States are free of M. arthritidis and conventionally maintained rats are rarely M. arthritidis antibody positive. The organism has been isolated, typically without pathological implications, in various nonarticular sites such as the submaxillary gland (Klieneberger, 1938); the oropharynx; middle ear; lung, usually with coinfection by M. pulmonis or other bacteria (Cole et al., 1967; Stewart and Buck, 1975); the large intestine; lower female genital tract; upper male genital tract; and the Harderian gland (Cox et al., 1988). Experimental inoculation in athymic nude rats and euthymic (rnu/þ) littermates resulted in a more severe arthritis and progressive disease leading to deaths in the athymic (rnu/rnu) rats (Binder et al., 1993). Naturally occurring M. arthritidis-induced arthritis has been rarely reported in rats. Nearly all the information associated with M. arthritidis-induced arthritis reflects experimental infections; however, the clinical signs seen in both naturally occurring and induced

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arthritis are similar. Typically, multiple joints are swollen, inflamed, and tender, with rats tending to avoid weightbearing on the affected limbs. The articular and paraarticular tissues are swollen, and the synovial cavity contains a thick, purulent exudate. After intravenous inoculation of M. arthritidis,polyarthritis becomes evident in 2e9 ays, with the arthritis resolving in about 2 months. Mycoplasma can be easily recovered during the peak stage of the disease but not after 6e8 weeks. Microscopically, there is an intense polymorphonuclear infiltration of the synovial space and membrane and of the surrounding articular tissue. This is followed by periostialosteoneogenesis and destruction of articular cartilage (Cole and Ward, 1979). Hindlimb paralysis may follow perineural inflammation and edema that compresses nerve roots within the spinal foramina (Cassell et al., 1979). The rat has been widely used to study the immunologically based pathogenesis of articular disease induced by intravenous inoculation of M. arthritidis (Cole and Ward, 1979). Although the mechanisms contributing to the pathogenesis of M. arthritidis are not well defined, of considerable interest is the M. arthritidis mitogen (MAM) superantigen, which is a potent T-cell mitogen and inducer of IFN-g for murine lymphocytes. Superantigens, as a class, can induce autoimmunity and act as potent immunomodulatory molecules. Although the MAM superantigen probably has a role in the arthritogenicity of M. arthritidis, all strains of the organism release MAM during lytic cell senescence (Cole and Atkin, 1991; Anh-Hue et al., 2002). A lysogenic bacteriophage, MAV1, is considered to be a virulence factor associated with arthritis in rats and mice because MAV1 lysogens are more arthritogenic than are nonlysogens. One of the MAV genes transcribed is thought to code for a membrane surface-exposed lipoprotein. This lipoprotein, MAA1, may mediate adherence of M. arthritidis to joint tissues and thus enhance its pathogenicity (Washburn et al., 2000; Anh-Hue et al., 2002). An excellent review of the pathogenic mechanisms involved in M. arthritidisinduced murine arthritis is presented by Washburn and Cole (2002). ELISA testing is the most frequent means to survey for M. arthritidis; however, because of one-way crossreacting antigens with M. pulmonis, immunofluorescence assay or a Western blot must be done to confirm if a positive ELISA serum sample reflects M. arthritidis infection. The effects of M. arthritidis on research are negligible compared with those of M. pulmonis. Naturally occurring latent infections have been shown to interfere with experimentally induced models of arthritis, and experimental infections have been shown to alter immunologic responsiveness (Cassell et al., 1986).

E. Mycoplasma haemomuris (Hemotropic Mycoplasma) Mycoplasma haemomuris, previously Haemobartonella muris, is one of four Mycoplasma spp. that have been transferred from the genera Haemobartonella and Eperythrozoon, the other three being Mycoplasma haemofelis, Mycoplasma haemosuis, and Mycoplasma wenyonii (Neimark et al., 2002; do Nascimento et al., 2012). M. haemomuris is an extracellular parasite of erythrocytes in rats and mice. Two genetically distinct clusters among strains of M. haemomuris have been demonstrated by analysis of the primary and secondary structures of the nucleotide sequence of the intergenic transcribed spacer region and the 16S rRNA gene. One cluster was isolated from mice and the second from black rats. The authors of the report conclude that this finding may support the hypothesis that the two clusters represent M. haemomuris subsp. musculi and M. haemomuris subsp. ratti (Sashida et al., 2013). Using Romanowsky stain, the organism appears as purple dots in blood smears. This Gramnegative organism is pleomorphic varying from a 350nm coccus to a 700-nm rod. It is an obligate parasite that is not cultivable outside the host (Lindsey et al., 1991). The blood-sucking rat louse Polyplax spinulosa is the vector necessary for transmission of M. haemomuris between rats. Today, because of the elimination of P. spinulosa from essentially all institutional resources housing rats, this mycoplasmal infection would very rarely be present in laboratory rats. Exceptions to this are colonies contaminated by wild or feral rats infected with both M. haemomuris and the vector. If the reticuloendothelial system is intact, natural infections of M. haemomuris are invariably latent. However, if the host has been splenectomized, there is a rapid increase in the number of organisms, leading to hemolytic episodes within 3e10 days, dyspnea, hemoglobinuria, and often death. Splenomegaly is essentially the only change seen upon gross examination of rats subsequent to natural infection with the agent. Red blood cell smears show M. haemomuris as coccoid and rod bodies arranged singly, and in chains or clusters on erythrocytes in splenectomized rats, but parasitemia would be infrequently detected in nonsplenectomized rats (Cassell et al., 1979). Transmission has been reported from contaminated blood, tumors, and other blood-laden biologic material (Cassell et al., 1979; Lindsey et al., 1991). Receipt of these biological materials from unreliable sources poses the greatest risk of infection to a naive group of recipient rats. Although transplacental transmission has been suggested, there is negligible evidence to support vertical transfer of the agent (Owen, 1982). Splenectomy, as a means to activate a latent infection, is used to detect the organism’s presence within a

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colony. Older rats develop more severe parasitemias than do young rats after splenectomy and, accordingly, should be used as subjects for activation. Positive identification of M. haemomuris in blood smears is achieved by use of Giemsa or Romanowsky stains; however, care must be taken not to confuse normal basophilic stippling of erythrocytes with the organism (Cassell et al., 1979). A specific and sensitive PCR test has been developed for the detection of M. haemomuris in mouse blood samples (Zhang and Rikihisa, 2002). The selected primers did not anneal with any gene sequences other than the 16S rRNA gene of M. haemomuris, including M. pulmonis, nor did they amplify closely related M. haemofelis and M. haemosuis. The most likely research complications associated with M. haemomuris would be through accidental transmission by contaminated blood, plasma, tumors, or other biological material. It is extremely important that investigators and veterinarians are aware that not all sources of biologic materials are derived from pathogen-free rats and mice, and that these materials could be contaminated with viral, bacterial, Mycoplasma,or ricketsial agents. M. haemomuris infection has been shown to affect transplantable tumor kinetics, reticuloendothelial function, erythrocyte half-life, IFN production, and host response to experimental infections with viral and protozoal agents (Sacks and Egdahl, 1960; Stansly, 1965; Cassell et al., 1979; Lindsey et al., 1991).

the brain, causing compression of neurons. Eight hours after inoculation, there are focal areas of spongiform degeneration, most prominently in the deep layers of the frontoparietal cortex and underlying white matter and in the molecular layer of the cerebrum (Thomas and Bitensky, 1966; Thomas et al., 1966).

F. Mycoplasma of Negligible Pathogenicity in the Rat

Dermatomycosis (dermatophytosis, ringworm, or favus) has long been associated with wild and laboratory rodents. Although the literature, particularly older reports, is rife with synonymy, the etiology of the majority of naturally occurring rodent ringworm is caused by T. mentagrophytes. This species, however, is one of the most polymorphic of the dermatophytes, and failure to recognize its range of forms has led to confusion in taxonomy. Two principal forms of the organism are recognized: (1) a zoophilic variant with granular colonial surface and red pigmentation named T. mentagrophytes var. T. mentagrophytes and (2) an anthropophilic form with white, fluffy colonial surface and no pigmentation designated T. mentagrophytes var. inter-digitalis (Ajello, 1974). Molecular approaches have been used to classify T. mentagrophytes substrains on the basis of DNA sequences (Makamura et al., 1998; Kim et al., 2001). Many reports, particularly in the mouse, refer to Trichophyton quinckeanum as a separate and distinct species, but most modern mycologists consider T. quinckeanum to be synonymous with T. mentagrophytes var.

1. Mycoplasma neurolyticum M. neurolyticum has been associated with conjunctivitis in mice. In the rat, there are no reports of naturally occurring disease induced by the organism, and there is essentially no evidence that it infects rats even as a commensal (Cassell et al., 1979). Most notably, the organism induces “rolling disease” in mice after intravenous, intraperitoneal, or intracerebral inoculation of the organism or cell-free culture filtrates (Findlay et al., 1938; Thomas and Bitensky, 1966). Rats, after intravenous inoculation of M. neurolyticum, develop neurological signs in less than an hour. Initially, animals become weak, ataxic, and prostrate; this is followed by clonic convulsive seizures and usually death in 3e4 h. In a few rats, there is a prolonged survival of several days. The pathogenicity of M. neurolyticum is caused by an exotoxin (Tully, 1969; Kaklamanis and Thomas, 1970), and its action induces severe distension of astrocytes and a generalized edema of

2. Mycoplasma volis This mycoplasma species was isolated from the respiratory tract of clinically normal prairie voles (Microtus ochrogaster). To test the possible pathogenicity of this newly defined species in rats, 10 Sprague-Dawley rats were inoculated intranasally with M. volis (EvansDavis et al., 1998). No clinical signs of infection were induced in any of the rats during the 6-week observation period, although all animals seroconverted to M. volis. The only histopathological changes were mild to severe hyperplasia of the bronchial-associated lymphoid tissue. M. volis was isolated from the nasopharynx of 9 rats and the lungs of 10 rats. The degree of peribronchial lymphoid hyperplasia was less at 6 weeks compared with that observed at 4 weeks after inoculation. This would tend to suggest that the rat is able to clear M. volis without long-term sequelae.

IV. MYCOTIC INFECTIONS A. Trichophyton mentagrophytes (Dermatomycosis)

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mentagrophytes (Ajello et al., 1968). Both variants may infect laboratory rodents. Much less frequently, several other dermatophytic species have been encountered in both wild and laboratory rodents (Feuerman et al., 1975; Kunstyr, 1980; Papini et al., 1997; Connole et al., 2001). Dermatomycosis is more common as a disease of laboratory mice (Parish and Craddock, 1931; Booth, 1952; Brown and Parker, 1957; Menges et al., 1957; Dolan et al., 1958; Mackenzie, 1961; Cetin et al., 1965; Davies and Shewell, 1965; Reith, 1968) and guinea pigs (Menges and Georg, 1956; Kaffka and Reith, 1960; Mohapatra et al., 1964; Otcenasek et al., 1974; Owens and Wagner, 1975; Pombier and Kirn, 1975; Kunstyr et al., 1980a,b) than of rats (Dolan et al., 1958; Dolan and Fendrick, 1959; Georg, 1960; Povar, 1965; Mizoguchi et al., 1986), as the volume of literature indicates. Periodic survey work indicates that T. mentagrophytes is not uncommon in wild rats (Smith et al., 1957; Georg, 1960; Thierman and Jeffries, 1980) and mice (Brown and Suter, 1969; Chmel et al., 1975), although, as will be pointed out later, the asymptomatic carrier state can be more common than is realized. The disease in rats may assume an epizootic form, with many of the animals showing lesions, or may be insidious and characterized by lesionless carriers. In a recently reported outbreak in Iraq, the prevalence of clinically apparent cases was 28% (Issa and Zangana, 2009). In both presentation modes, substantial hazard to human contacts is present; indeed, human infection by persons handling the animals is frequently the first indication that the infection is in the colony. Most human infections occur on the exposed, relatively hairless parts of the body, especially the hands and arm. As mentioned, the infection assumes variable form in rats and is thought to be influenced by a number of factors that include those directly bearing on susceptibility or resistance, for example, age, genetic constitution, immunologic competence, and phase of hair growth cycle, as well as other less understood factors. It is known that infection immunity in rats is thymus dependent (Green et al., 1983, 1987). Cortisone injections to test the hypothesis of immunocompetence experimentally failed to influence degree of infection compared with that in untreated guinea pigs (Fisher and Sher, 1972). The few reported epizootics with lesioned rats all occurred in nonutilized animals before experimentation (Dolan et al., 1958; Povar, 1965; Mizoguchi et al., 1986; Issa and Zangana, 2009). Lesions, when present, may occur in the skin of any area but are most common on the neck, back, and base of the tail. Lesions are not as classically described, that is, uniformly discoid with alopecia and raised margins, but rather may have a scurfy or erythematous papular/pustular appearance with irregular, patchy hair loss. Lesions on the tail (typically

seen in the mouse) were not seen in rats by Povar (1965) in the outbreak he described. Diagnosis of dermatophytosis is established by demonstration of fungal elements in skin scrapings and isolation of the causative organism by culture. Histopathology of affected skin is supportive in the attribution of lesion development to isolated dermatophytes if invasion of epidermal structures can be demonstrated. Histologic sections stained with Gridley fungus stain reveal fungal elements in the superficial epithelium and invasion of hair follicles. Secondary invasion of fungal lesions by bacteria with suppurative inflammation is commonly observed and is the cause of kerion-like lesions in both animals and humans. The differential diagnosis for dermatophytosis should also consider other causes of similarly appearing skin lesions, including staphylococcal ulcerative dermatitis, fighting bite trauma, hair chewing or barbering by cage mates, and ectoparasitic hypersensitivity (Kunstyr et al.,1980). Skin scrapings should be carefully taken from the lesion periphery, mounted in 10% potassium hydroxide under a Vaseline-ringed cover slip, and examined under the microscope immediately and again in 30 min. When present, septate mycelia are observed in squamous cells. Small spore (2e3 mm) ectothrix invasion of hairs, especially near the base, are seen in T. mentagrophytes infections. Scrapings, similarly collected, should be inoculated onto the surface of a suitable agar medium and cultivated aerobically at room temperature for at least 10 days before discarding as negative. Suitable media include dermatophyte test medium (DTM) with color indicators (Carroll, 1974) or Sabouraud’s medium with cycloheximide and chloramphenicol to inhibit nondermatophytic contaminants (Rosenthal and Fumari, 1957; Rebell and Taplin, 1970). Typical microscopic features of T. mentagrophytes (macroconidia, spiral coils) should be demonstrated (Rebell and Taplin, 1970). Use of the MALDI-TOF MS procedure has been described to identify species within the T. mentagrophytes complex (Packeu et al., 2013). MacKenzie’s hair brush technique is used to evaluate the incidence of lesionless, asymptomatic carrier rats (Mackenzie, 1963; Rosenthal and Wapnick, 1963; Papini et al., 1997). This technique may be used to screen sample groups of rats to establish their status as asymptomatic carriers of dermatophytes, although in the absence of suspicious lesions, scheduled health surveillance testing for dermatophytes is not recommended by FELASA (Nicklas et al., 2002). There is some evidence to indicate that this state may occur frequently in the rat (Dolan et al., 1958; Dolan and Fendrick, 1959; Gugnani et al., 1971; Balsari et al., 1981; Papini et al., 1997) although it should be pointed out that all of these data were obtained from conventionally bred rats housed in conventional caging and facilities. The organism is not

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known to traverse the placenta and has not been recovered from barrier-reared laboratory rats. Asymptomatic carriers are more commonly recognized in the guinea pig, mouse, and cat (Fuentes and Aboulafia, 1955; Fuentes et al., 1956; Menges et al., 1957; Dolan et al., 1958; Rosenthal and Wapnick, 1963; Gip and Martin, 1964; Feuerman et al., 1975). In this technique, the animal to be screened is held over an opened Petri dish of suitable agar medium and the hair brushed with a sterile surgical scrub brush so that hairs, flakes, and desquamated cellular debris fall directly down on the medium surface. The plate is incubated as described earlier. Eradication and control in research colonies usually entails destruction of the affected groups and sterilization or disinfection of equipment and environmental surfaces. A modified rederivation approach to eradication in a breeding colony was described by Mizoguchi et al. (1986). This program entailed removal of all rats from the colony, disinfection of the premises with formalin and sodium propionate, and subsequent restocking of the colony with weanlings from nonlesioned dams dipped in sodium propionate before reintroduction. Although the clinical efficacy of feeding griseofulvin to treat dermatomycosis has had mixed results in other laboratory species (Cetin et al., 1965; Pombier and Kirn, 1975), its effectiveness for this purpose has not been evaluated in rats.

B. Deep Mycoses The deep or systemic mycoses (fungal infections other than dermatomycosis) are extremely rare in rats; indeed, two comprehensive reviewsdone covering the period up to 1954 (Schwarz, 1954) and another to 1967 (Smith and Austwick, 1967)dfailed to document a single episode in unimpaired laboratory rats. Both asymptomatic and lesioned carriers of several fungal species have been encountered in wild R. norvegicus and other rat species, and they may be regarded as playing some role as natural reservoir hosts for certain organisms. The reviews just cited may be consulted for information related to wild rats. Aspergillosis has been reported in naturally occurring outbreaks of pulmonary (Singh and Chawla, 1974; Gupta, 1978; Hubbs et al., 1991) and nasal (Nyska and Kuttin, 1988; Rehm et al., 1988) infections of laboratory rats. Rat strains are known to differ in the immune mechanisms of resistance to experimental aspergillosis (Mirkov et al., 2015). In their review of systemic mycoses as experimental diseases in rat hosts, Smith and Austwick (1967) concluded that rats were quite refractory to most fungal infections; however, a number of reports have documented aspergillosis in cortisone-treated laboratory rats and mice as unanticipated outbreaks (Sagi

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and Lapis, 1956; Sidransky and Friedman, 1959; Sethi et al., 1964; Sandhu et al., 1970). Experimentally, lesions of aspergillosis were more severe in rats treated with cortisone than those immunosuppressed with immuran (Turner et al., 1976). That immunity is important in resistance is suggested both by the lethal aspergillosis induced experimentally in mice with antilymphocyte serum (Swenberg et al., 1969a,b) and by the protection induced by immunization (Corbel and Eades, 1977). In these reports, as in the natural outbreaks just cited, pulmonary aspergillosis presented as granulomatous pneumonia with miliary distribution in that organ primarily, although hematogenous distribution to other organs with capillary nets, for example, liver and kidney, was observed to a lesser degree. In the two reports of Aspergillus rhinitis, the lesions involved granulomatous inflammation of the maxilloturbinates. Because of the importance of invasive aspergillosis in humans with varying causes of immunodeficiency or impaired lung function (e.g., asthma or cystic fibrosis), laboratory rats have been extensively developed as models on which to base experiments exploring the pathogenesis, diagnosis, and treatment of this condition. Readers are referred to several recent reviews of this model system (Dagenais and Keller, 2009; Ahmad et al., 2014). Diagnosis of aspergillosis is established by cultural retrieval of the organism and supportive histopathologic lesions. PCR has been used especially to detect early pulmonary aspergillosis (Zhao et al., 2010). Sabouraud’s agar with bacterial inhibitors (chloramphenicol, penicillin, and streptomycin) is satisfactory for isolation, but care must be used to avoid modern media designed for isolation of dermatophytes (e.g., DTM), because the cycloheximide (Actidione) used as inhibitor will prevent growth by fungal “contaminants” such as Aspergillus. Isolated Aspergillus cultures should be speciated according to the criteria outlined by Austwick (1974), which emphasizes subculture on CzapekeDox medium for optimal growth of differential anatomic features useful in speciation. Granuloma formation is typical of pulmonary and upper respiratory aspergillosis in all species, including the rat. The lesions are seen microscopically to consist of macrophage and epithelioid cells centrally, as well as peripheral aggregation with both acute and chronic inflammatory cells, eosinophils, and giant cells of the Langhans type (Singh and Chawla, 1974). Congestion, microscopic hemorrhage, and septal thickening occur in surrounding parenchyma. Aspergillus is seen quite distinctly in material stained with hematoxylin and eosin, but Gridley’s fungus stain or GomorieGrocott should be used for careful study. The Aspergillus species may be differentiated in tissue from those causing phycomycosis by the more uniform and smaller hyphae (2.5 mm in diameter), regular branching at 45-degree

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angles, and prominent septae. In contrast, phycomycotic fungi in tissue have thicker hyphae (10 mm in diameter),irregular branches, and are nonseptate. Phycomycosis (or mucormycosis) is the general term used to describe tissue infection by nonseptate fungi of the genera Mucor, Absidia, and Rhizopus. The infection is quite rare in all species and has most often been reported in association with primary conditions lowering resistance or otherwise predisposing to phycomycosis, for example, diabetes mellitus, cortisone treatment, and immunosuppressive therapy. Mycotic encephalitis may be induced experimentally in mice by cortisone treatment (Swenberg et al., 1969a,b). Phycomycotic encephalitis has been reported in juvenile rats as a naturally occurring disease before experimentation (Rapp and McGrath, 1975; Moody et al., 1986). The lesions presented as purulent, necrotizing foci with acute inflammatory responses. Phycomycotic nonseptate hyphae up to 10e15 mm diameter were demonstrated with various stains. The investigators emphasized that the immaturity of these rats (2e4 weeks) was predisposing, because neither their dams nor older rats in the same room developed signs or lesions. Blastomycosis has also been encountered as a rare pulmonary infection of laboratory rats by (presumptive) Blastomyces dermatitidis (Chang et al., 2009). The diagnosis in this spontaneously hypertensive rat was based on histopathology and PCR.

C. Pneumocystis carinii (Pneumocystosis) Pneumocystis infection was originally described as a cause of pneumonia in laboratory guinea pigs by Chagas (1909), who mistook the cysts he saw as forms of Trypanosoma cruzi. Again, in 1911, he saw the cyst forms (which he reported as trypanosomes) in lung tissue from a human case of Chagas disease (Chagas, 1911). Shortly afterward, similar pneumonias in rats were described as caused by a new organism named P. carinii by Delanoe and Delanoe (1912). The organism was reported several years later in English laboratory mice (Porter, 1915). There are, however, few references to rodent pneumocystosis in the intervening years before the era of radiation and antiinflammatory (cortisone) research in the 1950s and 1960s (Frenkel et al., 1966). At that time, scientists began to encounter clinically overt Pneumocystis infections, principally in rats, as complications of research protocols now recognized as inducing a state of immunosuppression. Interestingly, most references to unanticipated, “spontaneous” C. piliforme and C. kutscheri infections of rats date to the same era, for the same reasons. Over the years, until the 1980s, informed opinion had wavered on the kingdom in which to place Pneumocystis,

but historically the organism was widely thought to be a protozoan. Uncertainty still surrounds details of the life cycle of Pneumocystis. The issue is complicated, and even the terms used to describe morphologic life forms of Pneumocystis derive from protozoology (see review by Chabe et al., 2011). The mature form is termed a cyst. Within the cyst, visible more or less depending on the stain and stage of maturation are eight intracystic bodies or “sporozoites.” When excystation takes place, the sporozoites enter a trophic stage. The vegetative, trophic forms in turn become precysts, crescent-shaped cysts, and finally cysts (Kim et al., 1972; Yoneda et al., 1982; Cushion et al., 1988). Cyst production involves a sexually reproductive phase (Cushion et al., 1997; Cushion, 2003). An ste3-like pheromone receptor has been identified in the organism (Smulian et al., 2001). Although the morphologic stage variants just described have been recognized microscopically, the lack of a continuous in vitro culture system for propagation of Pneumocystis has hampered an understanding of the progression of the life cycle and definition of the infective form. Pneumocystis may be carried for several passages in tissue culture, with an increase of up to 10- to 20-fold in numbers, but a continuous in vitro culture method has so far proven elusive (Bartlett et al., 1979; Cushion and Walzer, 1984a,b; Cushion et al., 1985; Cushion and Ebbets, 1990; Aliouat et al., 1995; Beck et al., 1996; Atzori et al., 1998). All phases take place in the mammalian lung, although tracing methods using PCR commonly detect Pneumocystis DNA in extrapulmonic locations, for example, bone marrow, liver, blood stream (antigenemia), and spleen (Reddy and Zammit, 1991; Schluger et al., 1991; Chary-Reddy and Graves, 1996; Rabodonirina et al., 1997). Lesions in spontaneous rodent Pneumocystis pneumonia (PCP) are usually confined to the lung, but there are reports of scid mice developing extrapulmonic foci in the heart and spleen (Reddy and Zammit, 1991). The cyst walls stain best with GMS, PAS, or toluidine blue O stains (Thompson et al., 1982; Sundberg et al., 1989). Giemsa stain does not stain the cyst wall. The best single stain for diagnostic visualization of Pneumocystis forms in concentrated smears of lung homogenates or impression smears of cut lung surface appears to be the Diff-Quik and similar stains because all stages are stained. Although several lines of evidence, for example, histologic tinctorial qualities (argyrophilia), were consistent with relatedness to fungi (Walkeden, 1990), antifungal drugs were shown to be ineffective against PCP, although Pneumocystis has since been shown, in vitro, to be susceptible to some antifungals (Kaneshiro et al., 2000). Conversely, drugs with confirmed activity against Pneumocystis, for example, trimethoprim-sulfamethoxazole, fansidar, and pentamidine, have an antiprotozoan spectrum.

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Molecular and biochemical analyses have established that Pneumocystis is a genus of unusual eukaryotic single-celled and genetically complex fungi, an ascomycete but lacking in ergosterol (Edman et al., 1988; Cushion, 1993; Stringer et al., 1992, 1993, 2002). Pneumocystis is classified in Phylum Ascomyota, Class Pneumocystidomycetes, Order Pneumocystidales, Family Pneumocystidaceae, Genus Pneumocystis. The nomenclature for species of concern in laboratory animals is listed in Table 12.2 (adapted from Cushion et al., 2004; Aliouat-Denis et al., 2008; Weisbroth et al., 2006). Other species are recognized for the dog, pig, and ferret (Chabe et al., 2011). A word may be in order about nomenclature and terminology. The preferred clinical terminology for PCPs has shifted somewhat. The acronym PcP for Pneumocystis carinii pneumonia has replaced PCP, although the latter continues in common human clinical usage even though Pneumocystis jirovecii is the agent causing human pneumocystosis. Similarly, the acronym for idiopathic interstitial pneumonia, IIP (see later), as a descriptor of lesions for “rat respiratory virus” infection has been discontinued because the pneumonia is now known to be caused by P. carinii and is no longer idiopathic. Confusingly, the acronym IIP may also be seen meaning infectious interstitial pneumonia. Isolates of Pneumocystis from rats are taxonomically quite complex. A trinomial system has been adopted (Wakefield, 1998; Cushion, 1998) in which the various strains of Pneumocystis have been designated as special forms (formaespeciales, or f. sp.; Table 12.2). At least five species of rat Pneumocystis have been delineated by DNA analysis, but only two of them have occurred in laboratory rats, e.g., P. carinii and Pneumocystis wakefieldiae (formerly, P. carinii f. sp. ratti) (Cushion et al., 2004). The binomial P. carinii is reserved for the more common Pneumocystis species occurring in rats and the former designation P. carinii f. sp. carinii is now shortened (Stringer et al., 2001). Three additional species have been detected in wild rats, namely, P. carinii f. sp. rattus-secundi, P. carinii f. sp. rattus-tertii, and P. carinii f. sp. rattus-quarti (Cushion et al., 1993; Palmer et al., 2000). The forms occurring in laboratory rats (i.e., P. carinii and P. wakefieldiae) may be discriminated by PCR (Palmer et al., 1999, 2000; Schaffzin et al., 1999; Nahimana et al., 2001) or by immunoblotting (Vasquez et al., 1996), and investigators should do so because the two species vary in responses to drug therapy trials, the life cycles differ, and the use of mixed isolates from lung preparations (not uncommon) could lead to a confusing mixture of gene sequences in gene libraries. At present, P. carinii is believed to be the most prevalent strain in commercial rat producer stocks (Icenhour et al., 2001), although coinfection by more than one special form in the same rat is not uncommon (Cushion et al.,

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1993; Nahimana et al., 2001). The name given to the mouse strain is P. murina (Keely et al., 2004) and to the species occurring in rabbits Pneumocystis oryctolagi (Dei-Cas et al., 2006; Soulez et al., 1989). Animals, especially rats and mice and to a lesser degree rabbits and ferrets, have been an indispensable element of Pneumocystis research dating from Frenkel’s 1966 paper (Dei-Cas et al., 1998). The histologic features of pulmonary PcP in animals are nearly identical to those of the disease in humans (see description and illustrations in Percy and Barthold, 2001; Barthold et al.,2016). Animal models have provided the basis for most of our current concepts of the epizootiology, taxonomy, diagnosis, genetics, and therapy of pneumocystosis (Hughes, 1989; Walzer, 1991). In the past, rats and mice from conventional commercial breeding colonies of immunocompetent stocks have acted as convenient but erratic and unreliable sources of carrier Pneumocystis infections. The term “conventional” describes a rodent population in which one or more murine viruses cycle as enzootic infections and which may also be infected with a range of helminth, protozoan, arthropod, bacterial, and fungal pathogens. The pathogen load is emblematic of unshielded husbandry environments and the correspondingly high likelihood that such populations also inapparently carry Pneumocystis. Investigators needing Pneumocystis-infected rodents for research programs favored procurement from such colonies in the past because it was assumed they were likely to be “latent” carriers (Russell and McGinley, 1991) although it has since been shown that Pneumocystis infection dynamics are quite independent of murine virus status (Pesanti and Shanley, 1984; Cushion and Linke, 1991). After arrival at the user institution, these animals would be deliberately subjected to an immunosuppressive protocol that would culminate, after an inductive phase of about 4e8 weeks, with development of severe lesions of PcP with abundant Pneumocystis. Faced with impending disappearance of conventional Pneumocystis carrier sources as commercial rodent breeders mounted intensive rederivation programs to eradicate murine viruses (Bartlett et al., 1987), investigators have turned to the more reliable method of inducing PcP in pathogen-free rodents. In this procedure, animals are first rendered susceptible to overt infection by immunosuppressive treatments and then infected by intranasal or intratracheal installation of Pneumocystis inocula (Bartlett et al., 1987, 1988, 1990, 1991; Shellito et al., 1990; Boylan and Current, 1992) or by short-term housing with infected seed rats or mice (McFadden et al., 1991). More recent models of pneumocystosis include the athymic (nude) rat and athymic (nude) and scid mice, which offer physiologic analogs comparable to the HIV-infected human with PcP. At present, athymic rodents that do not require further immunosuppression are used as the preferred

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medium for propagation of Pneumocystis. Neonatal rabbits commonly develop transient lesions of pneumocystosis, without immunosuppressive treatment, but these infections resolve naturally as immunocompetence develops in the weeks after birth (Soulez et al., 1989). Genetic studies with Pneumocystis isolates derived from various mammalian hosts have shown that these organisms are quite different. The accumulating results of DNA studies directed to taxonomic analysis have established P. jirovecii and P. carinii as separate species. Based on genetic divergence studies (Stringer et al., 1993) and infective isolation for its human host, the human strains of this organism were designated P. jirovecii (Stringer et al., 2002), as earlier suggested by Frenkel (1976). Although DNA sequence polymorphism has been shown with human Pneumocystis isolates (Lee et al., 1993), suggesting that numerous strains of P. jirovecii exist, this diversity has been thought insufficient to designate either special forms of P. jirovecii or additional human species (Wakefield, 1998; Stringer, 2002). P. jirovecii has not been found in the lungs of any other mammal, including nonhuman primates (Wakefield et al., 1990; Stringer, 2002), nor may it establish productive infection even in scid mice (Durand-Joly et al., 2002). Accordingly, it has been concluded that humans do not carry rodent Pneumocystis and, conversely, that humans do not contract pneumocystosis as a zoonosis from animal contacts. Experience from several lines of inquiry supports the concept that Pneumocystis strains have diverged in genetic heterogeneity, progressing to speciation according to homology with a given mammalian host species (Shah et al., 1996; Stringer, 1993). Infectivity studies have demonstrated that heterologous hosts, even if immunodeficient, are refractory to replication of Pneumocystis derived from other host species (Gigliotti et al., 1993; Aliouat et al., 1994). Similarly, the ultrastructural morphology and isoenzyme diversity among Pneumocystis isolates from different host species are host specific and support evidence of genetic diversity demonstrated by molecular analysis (Mazars et al., 1997; Nielsen et al., 1998). For this reason, under natural circumstances, rats should not be regarded as vectors of Pneumocystis infections for other rodent species, and conversely other rodent species, including mice, are not carriers of Pneumocystis infections for rats. The common denominator of immunodeficiency that underlies PcP has been linked with the common occurrence of intercurrent or “spontaneous” PcP in immunodeficient (athymic and scid) mutant and transgenic rat and mouse stocks, with induced rodent models for human PCP, and of course with human PCP, itself a consequence of impaired immunocompetence resulting from HIV infection. Progressive wasting (emaciation or cachexia), cough, dyspnea, and cyanosis are clinical signs

of PcP in rats. Clinically overt signs of pneumocystosis occur in rats and mice under the following circumstances: 1. Immunologically competent rodents that have been rendered immunodeficient as a consequence of experimental treatments in which PcP develops as an unplanned complication of a research protocol, as cited earlier during earlier eras of radiation and antiinflammatory research. Such animals may have been infected at the source before arrival at the user institution or first become exposed to same host species Pneumocystis shedders at the institution and infected after arrival. 2. Unplanned outbreaks in immunologically deficient mutants lacking cell-mediated immune capability and T-cell-assisted antibody responses. Such mutants include the athymic (nude) mouse and rat, the beige triple-deficient mouse, the scid mouse, and a range of similarly affected heritably immunodeficient mutant and transgenic rodent models (Veda et al., 1977; Van Hooft et al., 1986; Weir et al., 1986; Walzer et al., 1989; Sundberg et al., 1989; Gordon et al., 1992; Deerberg et al., 1993; Furuta et al., 1993; Percy and Barta, 1993; Pohlmeyer and Deerberg, 1993). These animals need only adequate exposure to Pneumocystis to initiate development of PcP (Soulez et al., 1991). On this basis, athymic mice have been used in mouse colonies as sentinels for detection of Pneumocystis (Serikawa et al., 1991). The clinical course of PcP in immunodeficient animals may be fulminant but is more likely to be chronic. The microscopic qualities of the lesions that develop in immunodeficient mutants are the same as in immunocompetent rodents that develop PcP as a result of immunosuppressive treatments. There is no support in the literature for the view that immunodeficient mutants may carry Pneumocystis in a latent or clinically silent form, and further it is established that immunosuppressive treatments of athymic mutants do not increase the incidence or severity of PcP during experimental induction (Veda et al., 1977; Walzer and Powell, 1982; Soulez et al., 1991; McFadden et al., 1991). 3. Immunologically competent carrier rats and mice rendered immunodeficient by chemical immunosuppressants, for example, corticosteroids or antimetabolites such as cyclophosphamide, as a planned process to develop PcP for some further experimental purpose; to propagate Pneumocystis for preparation of inocula or to provide antigen material; or to detect Pneumocystis carriers through the outcome of a diagnostic stress test. This circumstance is largely historical due to the common availability of heritably immunodeficient rodents for propagation of the agent and to more efficient diagnostic technology, e.g., PCR.

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Clinically overt pneumocystosis (PcP) should be viewed in all species as an unusual outcome requiring special circumstances of infection (immunodeficiency) as outlined earlier. More commonly, pneumocystosis occurs as a clinically inapparent infection of immunocompetent rodent stocks and strains that, because of its symptomalogic silence and difficulty of diagnostic detection (until recently), seemed in the recent past to have provided little incentive to commercial and institutional rodent breeders for eradication. Moreover, because of its clinical silence, it was not recognized, at the time, that pneumocystosis in immunocompetent rats would, nonetheless, be accompanied by pulmonary lesions (PcP). For example, until recently both FELASA and some commercial rodent breeders only included testing for Pneumocystis in health monitoring of immunodeficient breeding units (see Table 12.1. Indeed, in 2001, most immunocompetent rats from good commercial producers in the United States were detected as contaminated with DNA of P. carinii (Icenhour et al., 2001, 2002; Weisbroth et al., 1999a). In the late 1990s, circumstances were reported of idiopathic inflammatory lesions in the lungs of rats used in research (see references to this point cited in Livingston et al., 2011) or reported as a result of health monitoring by some commercial rodent breeders (PritchettCorning et al., 2009; Weisbroth et al., 2006) in animals known to be free of common murine viruses and BMM. The condition was shown to be infectious, but subclinical, to occur naturally in rats 6e20 weeks of age, and for lesion intensity to decrease as the rats progressed in age from adolescence to adult. The characteristic pulmonary lesions were given the descriptor “idiopathic interstitial pneumonia” or IIP to underscore the etiologic uncertainty. The putative agent was given the provisional designation “rat respiratory virus” and the acronym RRV (Albers and Clifford, 2003; Albers et al., 2009; Besselsen et al., 2008; Riley et al., 1997, 1999). Since that time diagnostic investigation has determined that the IIP lesions attributed to RRV were actually the subclinical lesions of P. carinii infection in immunocompetent rat hosts (Henderson et al., 2012; Livingston et al., 2011; Kim et al., 2014). Subsequently, the designations RRV and IIP have been discontinued. It has become clear that the primary natural mode for transmission and maintenance of rodent Pneumocystis infection is by exposure via aerosol of susceptible immunocompetent hosts (Walzer et al., 1977; Hughes, 1982; Hughes et al., 1983) although exposure to dirty bedding or to direct contact with shedding carriers will also suffice to effect transmission (Henderson et al., 2012). On the other hand, even germ-free rats were shown to be insusceptible to oral dosing via the diet or by ingestion of infected lung (Hughes, 1982). In enzootically infected

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breeding colonies, neonates are born to variably immune dams, exposed within hours of birth (Icenhour et al., 2002), but, it may be assumed, are protected from productive infection for the first few weeks of life by colostral antibody. Thereafter they become susceptible and cage-to-cage transmission occurs via aerosol from infected shedder animals. In enzootically infected, closed, barriered breeding colonies, weanling rats (3e 4 weeks old [100e125 g]) can be envisioned as living in a cloud of Pneumocystis spores, with spore contamination, at least as denoted by PCR detection of Pneumocystis DNA, on all environmental surfaces (Icenhour et al., 2002), thus with ample opportunity for exposure. The prime infective (shedder) phase would thus appear to be the 6e12-week-old rat (200e250 g), which corresponds to the age at which most rats are procured and shipped to user institutions. Thereafter the immunologic response to these light (and clinically inapparent) infections supervenes, and Pneumocystis populations decline until eliminated by the host (An et al., 2003). Thus the infection in the colony would seem to be propagated by successive cohorts of the productively infected, spore-shedding 6e12-week age group, infectious only to the next cohort of 3e4-week-old rats that are in the process of losing their colostral immunity. It would appear that older rats may be immune to reinfection and younger rats protected by colostral antibody. As more has been learned about the pathogenesis of pneumocystosis in immunocompetent rat and mouse (and human) hosts, doubt has been cast as to whether the term “latent” ever actually applies to Pneumocystis infections. Even after PCP induced by immunosuppression, most rat hosts allowed to immunologically recover eliminate Pneumocystis from the lungs (Vargas et al., 1995), as do infected scid mice immunologically reconstituted with normative spleen cells (Chen et al., 1993). The basis for inadvertent introduction of asymptomatic and lightly infected shedder rodents into user institutions was outlined earlier as reflecting the enzootic infective cycle in breeding colonies in which it can be envisioned that essentially all rodents in the 4- to 6week to 12-week adolescent age group become infected for a brief period of time, after which they become immune noncarriers (An et al., 2003; Menotti et al., 2013). During this period (the shedder phase), they are infective hazards for other Pneumocystis-free rodents of the same species and have been a frequent cause of clinically overt infective “breaks” in immunodeficient stocks at user institutions (Dumoulin et al., 2000). This same infectivity model is believed to hold with humans as well, who become almost universally exposed and immune at a very young age (Stringer et al., 2002). In the same way, it is now understood that human PCP derives not from activation of endogenous “latent” infections

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but more probably from chance or nosocomial aerosol exposure to spores during a receptive phase of immunodeficiency (Chen et al., 1993; Vargas et al., 1995). Although the infective form of Pneumocystis in aerosols has still not been defined, there is ample evidence of Pneumocystis DNA detected by PCR of spore trap air samples (Wakefield, 1994,1996; Olsson et al., 1996; Menotti et al., 2013). Immunosuppression, as by HIV infection or preparation for organ transplantation in humans or by immunosuppressive treatment of immunocompetent rodents, can effectively override naturally acquired or experimental immunity and render even immune hosts susceptible to PCP (Shellito et al., 1990; Harmsen et al., 1995). Antibodies, whether derived actively after infection or passively by injection of immune sera, adequately protect against experimental homologous Pneumocystis infection (Gigliotti and Harmsen, 1997; Bartlett et al., 1998). Definitive diagnosis of PCP in rats continues to be based on morphologic criteria of consistent gross and microscopic lesions of lobar pneumonia, with microscopic correlates of interstitial pneumonia, lymphohistiocytic paravascular cuffing, hemorrhage, and alveolar distension with foamy, pinkish material in hematoxylin and eosin-stained material. These lesions are sufficiently characteristic to be regarded as pathognomonic. With use of special stains (e.g., GMS), multiple discoid blackish staining Pneumocystis cysts may be demonstrated in the alveoli. Smears of lung tissue prepared at necropsy may be stained with toluidine blue O or GMS to detect cyst forms of Pneumocystis in postmortem material. Morphologic diagnosis of PcP can be confirmed by PCR tests (Kitada et al., 1991; Schluger et al., 1992; O’Leary et al., 1995), and for this purpose, individual or pooled lung tissue, bronchioalveolar lavage, or oral swab samples may be used (Feldman et al., 1996; Weisbroth et al., 1999a). In considering the use of serology as a diagnostic screening tool, the many practical hurdles and potential sources of false-negative results impose important limitations to adoption of this method. It has been shown that the host species origin of Pneumocystis antigen determines the serologic specificity of anti-Pneumocystis antibodies (Walzer and Rutledge, 1980; Kovacs et al., 1989; Bauer et al., 1991, 1993). Although many investigators use serology to detect prior exposures to Pneumocystis (Peglow et al., 1990), rats are quite variable in the age when detectable levels of antibody appear (Walzer et al., 1987). Antibody levels in sera after homologous experimental infections, that is, rat origin Pneumocystis used to infect rats, have a much higher titer against homologous Pneumocystis antigen than do the same sera tested against Pneumocystis antigens derived from other host species. Scientists recognized as early as 1966 (Frenkel) that Pneumocystis antigens derived from either rats or

humans would fix complement only with serum from homologous hosts, although when tested by immunoblotting, sufficient cross-reactivity of human sera against rat trophozoite antigens has been shown to be diagnostically useful (Chatterton et al., 1999). A further potential cause of false negative could accrue to the point that, at least in rats, there are multiple species and special forms of P. carinii with an unestablished degree of crossreactivity. These problems combine to make unlikely the development of a universal antigen that would detect antibodies to Pneumocystis in all laboratory species, or even in just rats and mice. Rather, it would seem that homologous antigen/antibody systems would need to be used. Because of the practical issues related to the present inability for large-scale propagation of Pneumocystis, rat origin antigens are not commercially available and serodiagnosis is not commonly used for diagnostic screening. In their study confirming the identity of lesions previously attributable to RRV with those of PcP caused by P. carinii, Henderson et al. (2012) described use of an IFA test for rat sera using purified whole-cell homologous antigens that had been propagated in rnu (athymic) rats. Diagnosis of light and clinically inapparent rat infections, as is the case in immunocompetent rat stocks, in the past depended on use of a diagnostic stress test (for a review of procedural parameters used in conducting rodent stress tests, see Weisbroth, 1995). The traditional stress test required histologic demonstration of Pneumocystis in stained lung sections after use of chemical immunosuppressants and a low-protein diet during a 2- to 4-week induction period (Frenkel et al., 1966; Walzer et al., 1979, 1980; Milder et al., 1980). The test was also used, when negative, to provide evidence that the institutional or commercial source of the animals in the test were free of contamination by Pneumocystis. Stress tests are lengthy, cumbersome, expensive, and have been shown to be no more effective in detection of inapparent infection than are the same rats tested by PCR at the start of the procedure before induction (Weisbroth et al., 1999a,b). For this reason, the stress test as a screening method for the diagnosis of Pneumocystis infection has largely been discarded in favor of PCR. As mentioned earlier, for this purpose individual or pooled lung tissue, bronchioalveolar lavage, or oral swab samples can be used for screening purposes (Feldman et al., 1996; Icenhour et al., 2001; Weisbroth et al., 1999a). Because of the sensitivity of PCR, immunocompetent (but Pneumocystis-free) rats may be used as sentinels to monitor Pneumocystis status of given rat environments (Icenhour et al., 2001). There are very few options for the control and eradication of Pneumocystis from rat populations. Due almost entirely to the role of rats as prime models for exploration of treatment modalities for human PCP, there is an

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extensive body of experience in the treatment of clinically apparent PCP in rats. The generality is that although a fair number of treatments, including trimethoprim-sulfamethoxazole, dapsone, or pentamidine, as well as an impressive array of newer therapeutics (beyond the scope of this text for review), have been shown effective in amelioration of clinical signs (and lung counts), none have been shown effective in eradication of Pneumocystis (Hughes, 1979, 1988). For that reason, treatment options would appear limited to those few instances in which the life of a small population of immunodeficient animals needed to be prolonged for some purpose. Gnotobiotic rederivation into isolators followed by introduction to a rodent barrier stringently managed for exclusion of rodent pathogens is recommended as effective for eradication and restart of rat populations free of Pneumocystis. There is no convincing evidence that P. carinii can traverse the rodent placenta to effect vertical transmission moreover, there is a good deal of experience demonstrating efficacy of gnotobiotic rederivation (Wagner, 1985; Ito et al., 1991).

D. Encephalitozoon cuniculi (Microsporidiosis) E. cuniculi is an obligate, intracellular, eukaryotic organism within the Kingdom Fungi and Phylum Microsporidia. Microsporidia have an extremely broad hostrange, including insects, invertebrates, fish, birds, and mammals, including humans. Interestingly, microsporidia were first described in 1870 with the identification of Nosema bombyscis as the pathogen causing the disease pebrine in silkworms (Pasteur, 1870). Because of its low host specificity and environmentally resistant spores, animal to human infection through water and food contaminated by animal feces or urine, zoonotic infections can easily occur (Kotkova et al., 2013). The organism is of most concern in conventionally raised rabbits, particularly pet rabbits that usually are subclinically infected (Desoubeaux et al., 2017). Accordingly, individuals who are immunocompromised should use considerable caution if handling rabbits or other species that may be infected with the organism. In the 1960s E. cuniculi was reclassified as Nosema cuniculi, but again renamed E. cuniculi in the 1970s. Until fairly recently, and at the time of publication of the second edition of this text, microsporidia were considered to be protozoan organisms. Contributing to reclassification as Fungi are genetic and molecular characteristics, including closed mitosis, possession of a mitochondrial heat shock protein, a- and b-tubulins, and the presence of chitin and trehalose in spores, all components of fungi (Wasson and Peper, 2000). Using the availability of six complete microsporidian genomes, data strongly supportmicrosporidia as the earliest-diverging clade of sequenced fungi (Capella-Gutierrz at al., 2012). The E. cuniculi

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genome was the first complete microsporidian genome (2.9 Mbp) sequenced (Katinka et al., 2001; Keeling and Slamovits, 2004), and later Encephalitozoon intestinalis with a genome of 2.3 Mbp. Both species reflect that microsporidian nuclear genomes are the most reduced and compacted of any eukaryotic cell (Corradi et al., 2010). Although today there is general agreement that microsporidia are Fungi, there is still conflicting evidence that point to microsporidia having a protistan origin (Choi and Kim, 2017). Microsporidia typically induce latent infections in immunocompetent hosts; however, they cause severe and often lethal infections in immunocompromised hosts. Immunity to infection is a function of T-cells and dependent upon IFN-gand IL-12 (Han and Weiss, 2017). Although very infrequent in immunocompetent rats, granulomatous, nonsuppurative inflammatory lesions have been described within the brain, spinal cord, and, less often, the kidney (Majeed and Zubaidy, 1982). Unlike the rat, infection in the rabbit is characterized by granulomatous inflammation to a greater extent in other organs in addition to the brain, and subsequently a greater prevalence of clinical disease. The typical histopathological lesions and observation of the organisms within the lesions have historically been the means by which most animals have been shown to be infected. Spores are oval, 1.5  2.5 microns, staining poorly with hematoxylin and eosin and somewhat better with Giemsa. Conversely, Toxoplasma gondii stains well with hematoxylin and eosin. E. cuniculi spores are Gram positive, variably acidfast with a PAS-positive granule at the polar cap, stain dark magenta or purple with Goodpasture’s, and are argyrophilic. There is no cyst wall (Shadduck and Pakes, 1971). Serological testing for antibodies to the organism is accomplished by ELISA, immunofluorescence, and Western blot assays (Desoubeaux et al., 2017). Genomic DNA may be detected by PCR of tissues and fluids believed to be infected with the organism. Infection of the host cell involves a coiled-polar-tube organelle within the spore that everts and injects the spore cytoplasm into the host cell. After replication of intermediate forms, the host cell becomes distended, ruptures, and spores are released into the environment (i.e., intestinal tract, kidneys of host) (Han and Weiss, 2017). Vertical transmission has not been demonstrated in rats. In immunocompetent BALB/c mice, it has been shown after oral administration that E. cuniculi remains latent for extended periods and infected organs serve as sources of microsporidial spores. Upon immunosuppression, reactivation, and redissemination of E. cuniculi, infection occurs leading to clinical disease and lethality (Kotkova et al., 2013). Until the 1970s, before widespread availability of cesarean-derived and barrier-maintained rats and

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mice, the reported prevalence of E. cuniculi ranged from 20% to 50%. Attwood and Sutton (1965) reported E. cuniculi-induced cerebral granulomas scattered widely in the brains of 76 of 352 laboratory rats examined for a drug toxicology study. In rats and mice and particularly in rabbits, in which the prevalence was 50%e70%, the organism was considered to be responsible for interfering with the interpretation of much experimental data, primarily to its unrecognized presence in these animal hosts. Its latency also complicated outcomes of transplantable tumor experiments such as the Yoshida rat ascites tumor into recipient rats (Shadduck and Pakes, 1971). Today, the prevalence of E. cuniculi in laboratory rats is very low. A large-scale pathogen screening effort, in which over 22,000 serological samples from laboratory rats were tested, revealed a prevalence of 0.08% (North America) and 1.55% (Europe) for the organism. The authors of the report, however, note that this assay is more prone to false positives than most viral assays (Pritchett-Corning et al., 2009). A 2017 report (Marx et al.) of adventitial infections in mice and rats at 63 research facilities in the United States indicated that E. cuniculi was among infectious agents that 75%e95% of institutions excluded from their colonies. Over a 2-year period, an outbreak was reported from only one of these research facilities. The 2014 FELASA report lists E. cuniculi as an organism for which colony monitoring is optional and to be pursued if there is a specific need (Mahler et al., 2014).

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FURTHER READING

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