Microscopic and molecular identification of hemotropic mycoplasmas in South American coatis (Nasua nasua)

Accepted Manuscript Title: Microscopic and molecular identification of hemotropic mycoplasmas in South American coatis (Nasua nasua) Authors: Michelle P. Cubilla, Leonilda C. Santos, Wanderlei de Moraes, Zalmir S. Cubas, Christian M. Leutenegger, Marko Estrada, LeAnn L. Lindsay, Edvaldo S. Trindade, C´elia Regina C. Franco, Rafael F.C. Vieira, Alexander W. Biondo, Jane E. Sykes PII: DOI: Reference:

S0147-9571(17)30045-0 http://dx.doi.org/doi:10.1016/j.cimid.2017.06.004 CIMID 1146

To appear in: Received date: Revised date: Accepted date:

2-8-2016 30-5-2017 3-6-2017

Please cite this article as: Cubilla Michelle P, Santos Leonilda C, de Moraes Wanderlei, Cubas Zalmir S, Leutenegger Christian M, Estrada Marko, Lindsay LeAnn L, Trindade Edvaldo S, Franco C´elia Regina C, Vieira Rafael FC, Biondo Alexander W, Sykes Jane E.Microscopic and molecular identification of hemotropic mycoplasmas in South American coatis (Nasua nasua).Comparative Immunology, Microbiology and Infectious Diseases http://dx.doi.org/10.1016/j.cimid.2017.06.004 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

1

Microscopic and molecular identification of hemotropic mycoplasmas in South American coatis (Nasua nasua)

Michelle P. Cubillaa,b,*, Leonilda C. Santosc, Wanderlei de Moraesb,c, Zalmir S. Cubasb, Christian M. Leuteneggerd, Marko Estradad, LeAnn L. Lindsaye, Edvaldo S. Trindadea, Célia Regina C. Francoa, Rafael F.C. Vieirac, Alexander W. Biondo a,c, Jane E. Sykese

a

Department of Cell and Molecular Biology, Universidade Federal do Parana, Av. Cel. Francisco

H. dos Santos, s/n., Curitiba, PR 81531-980, Brazil b

Bela Vista Biological Sanctuary, Itaipu Binacional, R. Teresina, 62, Foz do Iguacu, PR 85866-

900, Brazil c

Department of Veterinary Medicine, Universidade Federal do Parana, R. dos Funcionarios,

1540, Curitiba, PR 80035-050, Brazil d

IDEXX Laboratories Inc., 2825 KOVR Drive, West Sacramento, CA 95605, USA

e

Department of Medicine & Epidemiology, School of Veterinary Medicine, University of

California, 2108 Tupper Hall, Davis, CA 95616, USA

*Corresponding author. E-mail address: [email protected] (M.P. Cubilla).

2

HIGHLIGHTS 

Hemoplasma was detected in Nasua nasua by three different techniques for microscopy.



Sequencing confirmed organisms resembling M. haemofelis and Mycoplasma sp.



Potential risk of transmission to human beings, domestic and wild animals.

Abstract Hemoplasmas were detected in two apparently healthy captive South American coatis (Nasua nasua) from southern Brazil during an investigation for vector-borne pathogens. Blood was subjected to packed cell volume (PCV) determination, a commercial real-time PCR panel for the detection of Anaplasma spp., Babesia spp., Bartonella spp., Hepatozoon spp., Leishmania spp., Mycoplasma haemofelis, ‘Candidatus Mycoplasma turicensis’, ‘Candidatus Mycoplasma haemominutum’, Neorickettsia risticii, Rickettsia rickettsii and Leptospira spp., and a panhemoplasma conventional PCR assay. PCV was normal, but both coatis tested positive for hemoplasmas and negative for all the remaining pathogens tested. Using different techniques for microscopy (light, confocal or SEM), structures compatible with hemoplasmas were identified. Sequencing of the 16S rRNA gene identified an organism resembling Mycoplasma haemofelis and another hemotropic Mycoplasma sp., with a sequence identity of 96.8% to a Mycoplasma sp. previously detected in capybaras. Keywords: Hemoplasma; Vector-borne disease; Wild mammal; Mycoplasma haemofelis

1. Introduction

3

South American coatis (Nasua nasua, Carnivora, Procyonidae) are gregarious, omnivorous and widely distributed mammals in South America, feeding mostly on invertebrates and fruits, but also on small vertebrates and animal carcasses [1]. In addition, foraging and grooming behavior may lead to association and intimate contact with other animal species, which may expose coatis to ticks [2], increasing the likelihood of vector-borne or blood-borne infections. Vector-borne pathogens have been previously reported in South American [3–6] and white-nosed coatis (Nasua narica) [7], which may be of public health concern due to their synanthropic and fearless behavior toward peridomestic environments, human beings and domestic animals [4,7,8] with frequent contact and attacks [8]. In such scenarios, coatis may play a role as potential reservoir for vectors and pathogens. Accordingly, the aim of the present study was to survey health status and vector-borne pathogens in captive South American coatis.

2. Materials and methods The present study was approved by the Animal Ethics Committee at the Agricultural Sciences Campus of the Universidade Federal do Paraná (protocol number 047/2014), and was conducted under regulations of the Brazilian Institute of Environment and Renewable Natural Resources (IBAMA).

2.1 Study and samples During the annual health assessment at Bela Vista Biological Sanctuary, Itaipu Binational, Foz do Iguassu city, Parana State, southern Brazil, physical examination and dental prophylaxis were performed on 2 captive adult South American coatis, one female and one male,

4

using a specific protocol for chemical restraint (ketamine 12 mg/kg, midazolam 0.6 mg/kg and meperidine 4 mg/kg, intramuscularly). Blood samples were collected using sterile EDTA and sodium heparin-containing vacuum tubes (Vacutainer, Becton & Dickinson Co, Franklin Lakes, NJ, USA) and thin blood smears were made immediately after collection. Packed cell volume (PCV) was determined and remaining EDTA samples were stored at -20°C until molecular analysis.

2.2 Light microscopy Peripheral blood smears were prepared, stained using May-Grünwald-Giemsa stain and examined using light microscopy (BX51, Olympus, Tokyo, Japan) under 1,000X magnification. Additionally, blood smears were evaluated in a confocal image system (A1RSiMP, Nikon, Tokyo, Japan) using CFI Plan Apo Lambda 100X/1.45 Oil .13mm WD objective with a 476 nm laser excitation and 482/35 nm bandpass emission barrier filter (blue) or 488 nm laser excitation and 525/50 nm bandpass emission barrier filter (green). For differential interference contrast (DIC) images, cells were excited using 488 nm laser and emission was collected using 525/50 nm bandpass. The confocal and DIC images were obtained using sequential systems of each channel (blue, green or DIC). Images were visualized with a commercial imaging software (Nis Elements 4.20, Nikon, Tokyo, Japan) and edited for publication (GIMP 2.8.16, available at http://www.gimp.org/).

2.3 Scanning electron microscopy (SEM) For SEM, 500 µL of sodium heparin anticoagulated blood was washed 3 times with phosphate buffered saline (PBS) by centrifugation at 600 g for 5 min. The resulting pellet was

5

rinsed with 0.1 M sodium cacodylate buffer, pH 7.4, and fixed at room temperature with modified Karnovski’s fixative [9] (2% glutaraldehyde, 4% paraformaldehyde and 1 mM CaCl2 in 0.1 M sodium cacodylate buffer, pH 7.4) for 1 h. After washing 2 times with 0.1 M cacodylate buffer, pH 7.4, cells were post-fixed and contrasted with osmium tetroxide (1% OsO4 in 0.1 M sodium cacodylate buffer, pH 7.4) at room temperature for 1 h in the dark and then washed 3 times with the same buffer. The resuspended cells were spread onto coverslips, dehydrated with increasing concentrations of ethanol, dried by the CO2 critical-point method (CPD 030, Bal-Tec, Balzers, Liechtenstein), sputter-coated with gold (SCD 030, Balzers Union, Balzers, Liechtenstein) and examined with a scanning electron microscope (TESCAN VEGA3 LMU, Tescan Orsay Holding, Brno, Czech Republic) at the Center for Electron Microscopy of the Federal University of Parana (CME-UFPR, Curitiba, PR, Brazil).

2.4 DNA extraction, PCR assays and sequencing Genomic DNA was extracted from 100 µL of EDTA blood using a commercially available kit (Quick-gDNATM MiniPrep Kit, Zymo Research Corp., Orange, CA, USA), according to the manufacturer’s instructions. Negative control purifications using ultra-pure water were performed in parallel to monitor cross-contamination. Samples were first screened using a commercial real-time PCR panel in single reactions able to detect the DNA of Anaplasma spp., Babesia spp., Bartonella spp., Hepatozoon spp., Leishmania spp., Mycoplasma haemofelis (Mhf), ‘Candidatus Mycoplasma turicensis’ (CMt), ‘Candidatus Mycoplasma haemominutum’, Neorickettsia risticii, Rickettsia rickettsii and Leptospira spp. (IDEXX Laboratories Inc., Sacramento, CA, USA, test code 2870). A PCR assay for the host 18S rRNA gene (internal sample control) was used to determine DNA quantity and

6

quality in addition to positive (synthetic DNA, Ultramer, IDT Technologies, Coralville, Iowa, USA) and negative (RNAse-free water, Thermo Scientific, Waltham, MA, USA) controls. Primers and probes for the target genes were based on IDEXX’s proprietary real-time PCR oligonucleotides (IDEXX Laboratories, Westbrook, ME, USA). Real-time PCR assays were validated using established protocols. Amplification efficiency was calculated using the linear equation E = 10 1/-s-1 (s: slope) and required to be between 95 and 105%. Thereafter, the nearly complete 16S rRNA gene (approximately 1,450 bp) was amplified from blood samples using primers 8F (5`-AGAGTTTGATCCTGGCTCAG-3`) and 1492R (5`GGTTACCTTGTTACGACTT-3`) [10], in order to perform phylogenetic analysis. Each reaction (50 uL) contained 1X reaction buffer, 1.25 U AmpliTaq Gold polymerase (Applied Biosystems, Austin, TX, USA), 1.5 mM MgCl2, 0.4 µM each primer, 200 µM each dNTP and 10 µL DNA template. Cycling conditions were 95°C for 10 min, followed by 35 cycles of amplification (95°C for 1 min, 48°C for 1 min and 72°C for 2 min), and a final extension at 72°C for 5 min, using a Dyad Peltier thermocycler (MJ Research Inc., Waltham, MA, USA), as described previously [11]. Genomic DNA, previously extracted from a cat blood containing 'Ca. M. haemominutum' and ultrapure water (Applied Biosystems, Austin, TX, USA) were used as positive and negative controls, respectively. The PCR amplification products were visualized on a 2.5% agarose gel containing GelStar nucleic acid gel stain (Lonza, Rockland, ME, USA) using a UV transilluminator. The cPCR assay (DNA extraction, reaction mix preparation, amplification and products analysis) was conducted in physically separate areas to prevent contamination. The DNA fragments were purified from the agarose gel (GenElute TM Gel Extraction Kit, SigmaAldrich Co., St. Louis, MO, USA) and sequenced by automated capillary electrophoresis Sanger method (ABI Prism 3730 DNA Analyzer, Applied Biosystems, Foster City, CA, USA; College

7

of Biological Sciences UCDNA Davis Sequencing Facility, University of California, Davis, CA, USA) employing the same primers used for amplification reaction. The sequencing chromatograms were manually examined and edited using MEGA 6.06 [12]. Sense and antisense sequences of each sample were assembled to generate the consensus sequence (MEGA 6.06 [12]), and the final data were investigated for the presence of chimeric sequences by USEARCH 8.1.1756 [13]. In order to verify the identity percentage with other sequences, BLASTN [14] was used.

2.5 Nucleotide sequence accession numbers Hemoplasma nucleotide sequences amplified from the two coatis were deposited in the GenBank database under the accession numbers KU554425 and KU554426.

2.6 Phylogenetic analyses Sequences generated from the 16S rRNA gene (fragments of 1,329 bp and 488 bp) were aligned with sequences from GenBank database using MUSCLE [15] and manually edited on MEGA 6 [12]. Phylogenetic trees were constructed by Bayesian Inference (BI) using MrBayes 3.2.6 [16] and Maximum Likelihood (ML) method using GARLI 2.1 (http://www.molecularevolution.org/software/phylogenetics/garli) [17]. The best nucleotide substitution model was determined by jModeltest 2.1 [18] and was set, in both methods, to the general time-reversible model [19] with gamma distributed rate variation across sites, four rate categories and a proportion of invariable sites. For Bayesian inference, the Metropolis-coupled Markov chain Monte Carlo (MCMCMC) analysis was applied to estimate the posterior probability distribution of sequences. Four Markov

8

chains were run for 1,000,000 cycles and sampled every 100th generation with the temperature set to 0.5. The run stopped when the standard deviation of split frequencies was < 0.01. Maximum likelihood analyses were run including 1000 bootstrap replicates. All reconstructions were visualized with FigTree 1.4.2 (http://tree.bio.ed.ac.uk/software/figtree) and edited with Inkscape 0.91 (http://www.inkscape.org).

3. Results Male and female coatis had weight of 4.88 kg and 8.02 kg respectively. Ectoparasites were not found. Both animals were considered healthy following physical examination. The PCV from male and female coatis, 37% and 32%, respectively, were within the reference values for the species (29.8-38.2%) [20]. Light microscopy images of stained blood smears revealed small basophilic epierythrocytic structures of coccoid shape, individually attached and often more than one per erythrocyte, or freely on smears, in contrast to acidophilic erythrocytes (Fig. 1). Confocal images of the dye autofluorescence were obtained in different excitation and emission of light. The erythrocytes were revealed in green (Fig. 2C) and small structures with nucleic acid in blue (Fig. 2A and 2C). Then, DIC showed coccoid structures, measuring approximately 0.4 µm in diameter (Fig. 2B, arrows), that aligned with the nucleic acid image, as observed in the overlay of fluorescent and DIC images (Fig. 2D). Epicellular organisms were also observed by using SEM, with 0.2-0.4 µm in diameter (Fig. 3) and one organism demonstrated what appeared to be binary fission (Fig. 3B). These structures, revealed by different techniques of microscopy, were suggestive of hemotropic mycoplasmas. Using real-time PCR, both coati blood samples tested positive for Mhf and CMt, showing threshold cycle values of 35.32 (Mhf) and 33.73 (CMt) for the female coati and 30.52 (Mhf) and

9

39.56 (CMt) for the male coati. Samples tested negative for all the remaining pathogens. Amplification efficiencies were between 95 and 105%. Both coati blood samples tested positive for hemoplasmas using cPCR. The cPCR product from the female coati sample was sequenced in both forward and reverse directions and a consensus sequence of 1,329 bp was obtained (GenBank accession number KU554425). The cPCR product from the male coati sample was successfully sequenced only in the reverse direction and yielded a 488 bp fragment (GenBank accession number KU554426). A BLAST analysis of the sequence from the female coati (KU554425) revealed 96.8% and 95.9% identity with 16S rRNA gene sequences of Mycoplasma spp. amplified from the blood of Brazilian capybaras (Hydrochaeris hydrochaeris) (GenBank accession number FJ667773 and FJ667774, respectively); 93.0% (GenBank accession number KC863983) and 92.8% (GenBank accession number KJ739311) identity with 16S rRNA gene sequences of Mycoplasma spp. amplified from the blood of rodents from Hungary (Micromys minutus and Rattus norvegicus, respectively); and 91.3% with the 16S rRNA gene sequence of CMt amplified from a cat from Switzerland (GenBank accession number AY831867). A BLAST analysis of the 488 base pair sequence amplified from the male coati (KU554426) showed 100% identity with a 16S rRNA gene sequence of a Mycoplasma sp. amplified from the blood of raccoons (Procyon lotor) from USA (GenBank accession number KF743735) and 99.6% identity with a 16S rRNA gene sequence of Mhf amplified from a lion (Panthera leo) from Tanzania (GenBank accession number DQ825451). High levels of identity (99%) were also observed with many M. haemofelis 16S rRNA gene sequences amplified from domestic cats and other Felidae. The relationships of the sequences KU554425 and KU554426 amplified from the coatis with those from previously identified hemoplasmas are shown in Fig. 4. The phylogenetic

10

consensus trees obtained using BI and ML methods revealed the same phylogenetic relationships between the significantly supported clades. Both sequences were positioned in the haemofelis cluster. The sequence KU554425 from the female coati clustered as a basal sister clade to Mycoplasma spp. sequences detected in Brazilian capybaras (GenBank accession number FJ667773 and FJ667774), supported by a high BI posterior probability (100%) and ML bootstrap (100%) values. The sequence KU554426 from the male coati clustered in a well supported M. haemofelis / M. haemocanis 16S rRNA clade, species that have nearly identical 16S rRNA gene (>99% identity) [21], with ML bootstrap and BI posterior probability values of 100%.

4. Discussion The present study describes evidence of hemoplasma infection in South American coatis. A previous study detected a hemotropic Mycoplasma sp. from 20 white-nosed coatis from Costa Rica, which was most closely related to Mycoplasma haemolamae (82.8% identity) [7], but unfortunately, this 16S rRNA gene sequence was not available in the GenBank database for comparison. Herein, analyses of partial sequences of 16S rRNA gene identified two hemoplasma species infecting coatis. One Mycoplasma sp. (GenBank accession number KU554425) appeared to be most closely related to a hemotropic Mycoplasma sp. previously identified in capybaras from the same geographical region [22], with 96.8% and 95.9% identity (GenBank accession numbers FJ667773 and FJ667774, respectively). Although similarities less than 97% in the 16S rRNA gene sequences between two bacterial organisms suggest that they belong to different species, further molecular characterization using less conserved genes such as rpoB, 23S rRNA and ITS sequence are needed to confirm this hypothesis [23]. Phylogenetic analysis confirmed the close relationship of the female coati hemoplasma with the hemoplasmas

11

detected in capybaras, the sequence positioned as a sister clade to the capybara hemoplasmas. Coatis and capybaras differ in ecology, behavior and order (Carnivora and Rodentia, respectively); however, these animals are co-located geographically, and may be exposed to same vectors. Anemia was reported in some of the capybaras infected with hemoplasmas [22], but the female coati showed no clinical signs of illness and had a normal PCV. Although the real-time PCR assay yielded positive results for Mhf in both coatis, infection with this organism (or a closely-related organism) could only be confirmed by sequence analysis in the male coati. Phylogenetic analysis of the partial sequence available (GenBank accession number KU554426) showed 100% identity with Mycoplasma sp. previously found in the blood of North American raccoons, which are also members of Procyonidae family along with the South American coatis. Mycoplasma haemofelis has been considered the most pathogenic species of hemoplasma infecting cats and often associated with hemolytic anemia [24,25], but this was not observed in the male coati, which also had no clinical signs and a normal PCV. Since Mhf infection has been reported in an HIV-positive patient with positive cats and multiple cat scratches and bites [26], Mhf may be a less host-specific pathogen and therefore have a higher zoonotic potential. Alternatively, sequencing of additional genes may reveal that the hemoplasma may in fact be a different species to Mhf, as is the case for Mhf and Mycoplasma haemocanis, which share the same 16S rRNA gene sequence but have different RNase P gene sequences, different morphology, and differ in host species infected. Despite CMt was detected in both coatis using real time PCR, infection could not be confirmed by cPCR and sequencing. Although other vector-borne pathogens have been reported in coatis [3–7], and their role as reservoir of Trypanosoma cruzi [4] has been discussed, the coatis tested negative for all other vector-borne pathogens in the real-time PCR assay.

12

Hemoplasmas were detectable in the coati blood samples using three different microscopy techniques: conventional light microscopy, confocal microscopy, and SEM. With every technique, the microorganism appeared as small coccoid structures attached to the surface of the erythrocytes, as previously described for other hemoplasma species [11,27–31]. Confocal and SEM examinations of the female coati sample revealed epierythrocytic structures that approximated the size of small hemoplasma species like 'Ca. M. haemominutum' [27,28], CMt [28], 'Ca. M. haematoparvum' [11] and 'Ca. M. kahanei' [29]. In addition to molecular methods, confocal and electron microscopy have improved understanding of host-pathogen relationships because they provide greater morphological and morphometrical detail when compared with light microscopy [28,29,31,32]. Besides indirect transmission by arthropod vectors, direct transmission of hemoplasmas through aggressive contact among cats has been suggested [33–35]. Recently, sequences closely related to M. haemocanis/M. haemofelis and to Mycoplasma sp. detected in a capybara from Brazil were also detected in wild South American coatis from central-western Brazil [36], supporting the hypothesis that hemoplasmas could infect this species. In that study, ticks and fleas collected from the analyzed animals tested negative for Mycoplasma spp. [36]. Due to the close contact of coatis with other wild and domestic animals and human beings [4,7,8], their role in the transmission of hemoplasmas should be further investigated. The Bela Vista Biological Sanctuary is surrounded by houses and agricultural properties, and domestic cats and dogs from these properties were observed in the unity. Furthermore, the sanctuary is in a forest corridor and wild felids and other animals from local fauna were also observed in nearby areas, such as puma, jaguar, coatis and capybaras. This environment, together with the presence of vectors may facilitate cross-species infection.

13

5. Conclusion An organism that closely resembled Mycoplasma haemofelis as well as a Mycoplasma sp. most closely related to a hemoplasma from capybaras were identified in South American coatis. Special attention should be given to the potential risk of transmission to human beings, domestic animals and other wild animals.

Acknowledgements The authors acknowledge the Center for Electron Microscopy and to the Confocal and Conventional Fluorescence Microscopy Multi-user Laboratory at UFPR, for technical support and to the Coordination for the Improvement of Higher Education (CAPES) - Brazilian Ministry of Education, for the Post-graduate Support Program (PROAP). M. P. Cubilla was funded by the Brazilian National Council for Scientific and Technological Development (CNPq) – Brazil. The project was supported in part by IDEXX Laboratories.

References [1]

M.E. Gompper, D.M. Decker, Nasua nasua, Mamm. Species. 580 (1998) 1–9. doi:10.1126/science.95.2469.427-b.

[2]

I. Sazima, What coatis and mongooses have in common?, Biota Neotrop. 10 (2010) 457– 461. doi:10.1590/S1676-06032010000300040.

[3]

A.F.S.F. Rodrigues, E. Daemon, C.L. Massard, Morphological and morphometrical characterization of gametocytes of Hepatozoon procyonis Richards, 1961 (Protista, Apicomplexa) from a Brazilian wild procionid Nasua nasua and Procyon cancrivorus

14

(Carnivora, Procyonidae), Parasitol. Res. 100 (2007) 347–350. doi:10.1007/s00436-0060268-y. [4]

H.M. Herrera, C. V. Lisboa, A.P. Pinho, N. Olifiers, R.C. Bianchi, F.L. Rocha, G.M. Mourão, A.M. Jansen, The coati (Nasua nasua, Carnivora, Procyonidae) as a reservoir host for the main lineages of Trypanosoma cruzi in the Pantanal region, Brazil, Trans. R. Soc. Trop. Med. Hyg. 102 (2008) 1133–1139. doi:10.1016/j.trstmh.2008.04.041.

[5]

F.M. Alves, N. Olifiers, R. de C. Bianchi, A.C. Duarte, P.M.T. Cotias, P.S. D’Andrea, M.E. Gompper, G. de M. Mourão, H.M. Herrera, A.M. Jansen, Modulating variables of Trypanosoma cruzi and Trypanosoma evansi transmission in free-ranging coati (Nasua nasua) from the Brazilian Pantanal region, Vector Borne Zoonotic Dis. 11 (2011) 835– 841. doi:10.1089/vbz.2010.0096.

[6]

R. Lainson, R.R. Braga, a a De Souza, M.M. Pôvoa, E. a Ishikawa, F.T. Silveira, Leishmania (Viannia) shawi sp. n., a parasite of monkeys, sloths and procyonids in Amazonian Brazil., Ann. Parasitol. Hum. Comp. 64 (1989) 200–207.

[7]

L.R. Mehrkens, L.A. Shender, M.J. Yabsley, B.C. Shock, F.A. Chinchilla, J. Suarez, K.V.K. Gilardi, White-nosed coatis (Nasua narica) are a potential reservoir of Trypanosoma cruzi and other potentially zoonotic pathogens in Monteverde, Costa Rica, J. Wildl. Dis. 49 (2013) 1014–1018. doi:10.7589/2013-01-005.

[8]

G.C. Bittner, N.R. Hans, G. Hans Neto, M.O. Morais, G. Hans Filho, V. Haddad Jr, Coati (Nasua nasua) attacks on humans: case report, Wilderness Environ. Med. 21 (2010) 349– 352. doi:10.1016/j.wem.2010.09.005.

[9]

M. Karnovsky, A formaldehyde-glutaraldehyde fixative of righ osmolality for use in electron microscopy, J. Cell Biol. 27 (1965) 137.

15

[10] C. Pitulle, D.M. Citron, B. Bochner, R. Barbers, M.D. Appleman, Novel bacterium isolated from a lung transplant patient with cystic fibrosis., J. Clin. Microbiol. 37 (1999) 3851–5. [11] J.E. Sykes, L.M. Ball, N.L. Bailiff, M.M. Fry, “Candidatus Mycoplasma haematoparvum”, a novel small haemotropic mycoplasma from a dog, Int. J. Syst. Evol. Microbiol. 55 (2005) 27–30. doi:10.1099/ijs.0.02989-0. [12] K. Tamura, G. Stecher, D. Peterson, A. Filipski, S. Kumar, MEGA6: Molecular evolutionary genetics analysis version 6.0, Mol. Biol. Evol. 30 (2013) 2725–2729. doi:10.1093/molbev/mst197. [13] R.C. Edgar, Search and clustering orders of magnitude faster than BLAST, Bioinformatics. 26 (2010) 2460–2461. doi:10.1093/bioinformatics/btq461. [14] Z. Zhang, S. Schwartz, L. Wagner, W. Miller, A Greedy Algorithm for Aligning DNA Sequences, J. Comput. Biol. 7 (2000) 203–214. doi:10.1089/10665270050081478. [15] R.C. Edgar, MUSCLE: Multiple sequence alignment with high accuracy and high throughput, Nucleic Acids Res. 32 (2004) 1792–1797. doi:10.1093/nar/gkh340. [16] F. Ronquist, M. Teslenko, P. Van Der Mark, D.L. Ayres, A. Darling, S. Höhna, B. Larget, L. Liu, M.A. Suchard, J.P. Huelsenbeck, Mrbayes 3.2: Efficient bayesian phylogenetic inference and model choice across a large model space, Syst. Biol. 61 (2012) 539–542. doi:10.1093/sysbio/sys029. [17] A.L. Bazinet, D.J. Zwickl, M.P. Cummings, A gateway for phylogenetic analysis powered by grid computing featuring GARLI 2.0, Syst. Biol. 63 (2014) 812–818. doi:10.1093/sysbio/syu031. [18] D. Darriba, G.L. Taboada, R. Doallo, D. Posada, jModelTest 2: more models, new

16

heuristics and parallel computing, Nat. Methods. 9 (2012) 772–772. doi:10.1038/nmeth.2109. [19] C. Lanave, G. Preparata, C. Sacone, G. Serio, A new method for calculating evolutionary substitution rates, J. Mol. Evol. 20 (1984) 86–93. doi:10.1007/BF02101990. [20] A.S. Labate, A.L.V. Nunes, M. da S. Gomes, Order Carnivora, Family Procyonidae (Raccoons, Kinkajous), in: M.E. Fowler, Z.S. Cubas (Eds.), Biol. Med. Surg. South Am. Wild Anim., Iowa State University Press, Ames, Iowa, USA, 2008, pp. 317–322. doi:10.1002/9780470376980.ch28. [21] A.J. Birkenheuer, E.B. Breitschwerdt, A.R. Alleman, C. Pitulle, Differentiation of Haemobartonella canis and Mycoplasma haemofelis on the basis of comparative analysis of gene sequences, Am. J. Vet. Res. 63 (2002) 1385–1388. doi:10.2460/ajvr.2002.63.1385. [22] R.F.C. Vieira, M.B. Molento, L.C. dos Santos, W. Moraes, Z.S. Cubas, A.P. Santos, A.M.S. Guimaraes, A. Mohamed, I.R. Barros Filho, A.W. Biondo, J.B. Messick, Detection of a novel hemoplasma based on 16S rRNA gene DNA in captive and freeranging capybaras (Hydrochaeris hydrochaeris), Vet. Microbiol. 139 (2009) 410–413. doi:10.1016/j.vetmic.2009.06.018. [23] M. Drancourt, D. Raoult, Sequence-based identification of new bacteria: A proposition for creation of an orphan bacterium repository, J. Clin. Microbiol. 43 (2005) 4311–4315. doi:10.1128/JCM.43.9.4311-4315.2005. [24] S. Tasker, Haemotropic mycoplasmas. What’s their real significance in cats?, J. Feline Med. Surg. 12 (2010) 369–381. doi:10.1016/j.jfms.2010.03.011. [25] J.E. Sykes, Feline hemotropic mycoplasmas, J. Vet. Emerg. Crit. Care. 20 (2010) 62–69.

17

doi:10.1111/j.1476-4431.2009.00491.x. [26] A.P. Dos Santos, R.P. Dos Santos, A.W. Biondo, J.M. Dora, L.Z. Goldani, S.T. de Oliveira, A.M. de Sá Guimarães, J. Timenetsky, H.A. de Morais, F.H.D. González, J.B. Messick, Hemoplasma infection in HIV-positive patient, Brazil, Emerg. Infect. Dis. 14 (2008) 1922–1924. doi:10.3201/eid1412.080964. [27] J.E. Foley, N.C. Pedersen, “Candidatus Mycoplasma haemominutum”, a low virulence epierythrocytic parasite of cats, Int. J. Syst. Evol. Microbiol. 51 (2001) 815–817. doi:10.1099/00207713-51-3-815. [28] B. Willi, K. Museux, M. Novacco, E.M. Schraner, P. Wild, K. Groebel, U. Ziegler, G.A. Wolf-Jäckel, Y. Kessler, C. Geret, S. Tasker, H. Lutz, R. Hofmann-Lehmann, First morphological characterization of “Candidatus Mycoplasma turicensis” using electron microscopy, Vet. Microbiol. 149 (2011) 367–373. doi:10.1016/j.vetmic.2010.11.020. [29] H. Neimark, A. Barnaud, P. Gounon, J.C. Michel, H. Contamin, The putative haemobartonella that influences Plasmodium falciparum parasitaemia in squirrel monkeys is a haemotrophic mycoplasma, Microbes Infect. 4 (2002) 693–698. doi:10.1016/S12864579(02)01588-5. [30] Y. Maede, M. Sonoda, Studies on feline haemobartonellosis. III. Scanning electron microscopy of Haemobartonella felis, Japanese J. Vet. Sci. 37 (1975) 209–211. doi:http://doi.org/10.1292/jvms1939.37.209. [31] N.C. do Nascimento, A.P. dos Santos, Y. Chu, A.M.S. Guimaraes, A.N. Baird, A.B. Weil, J.B. Messick, Microscopy and genomic analysis of Mycoplasma parvum strain Indiana, Vet. Res. 45 (2014) 1–11. doi:10.1186/s13567-014-0086-7. [32] K. Groebel, K. Hoelzle, M.M. Wittenbrink, U. Ziegler, L.E. Hoelzle, Mycoplasma suis

18

invades porcine erythrocytes, Infect. Immun. 77 (2009) 576–584. doi:10.1128/IAI.0077308. [33] B. Willi, F.S. Boretti, M.L. Meli, M. V. Bernasconi, S. Casati, D. Hegglin, M. Puorger, H. Neimark, V. Cattori, N. Wengi, C.E. Reusch, H. Lutz, R. Hofmann-Lehmann, Real-time PCR investigation of potential vectors, reservoirs, and shedding patterns of feline hemotropic mycoplasmas, Appl. Environ. Microbiol. 73 (2007) 3798–3802. doi:10.1128/AEM.02977-06. [34] K. Museux, F.S. Boretti, B. Willi, B. Riond, K. Hoelzle, L.E. Hoelzle, M.M. Wittenbrink, S. Tasker, N. Wengi, C.E. Reusch, H. Lutz, R. Hofmann-Lehmann, In vivo transmission studies of “Candidatus Mycoplasma turicensis” in the domestic cat, Vet. Res. 40 (2009) 45. doi:10.1051/vetres/2009028. [35] M.R. Lappin, P. Dingman, J. Levy, J.R. Hawley, A. Riley, Detection of hemoplasma DNA on the gingiva and claw beds of naturally exposed cats, J. Vet. Intern. Med. 22 (2008) 779. doi:10.1111/j.1939-1676.2008.0103.x. [36] K.C.M. de Sousa, H.M. Herrera, C.T. Secato, A.D.V. Oliveira, F.M. Santos, F.L. Rocha, W.T.G. Barreto, G.C. Macedo, P.C.E. de Andrade Pinto, R.Z. Machado, M.T. Costa, M.R. André, Occurrence and molecular characterization of hemoplasmas in domestic dogs and wild mammals in a Brazilian wetland, Acta Trop. 171 (2017) 172-181. doi:10.1016/j.actatropica.2017.03.030.

19

Fig.1. Light microscopy images of May-Grünwald-Giemsa-stained blood smears from coatis, showing small basophilic structures attached to erythrocytes (arrows). A: Sample from female coati. B: Sample from male coati (1,000X). Bar = 10 µm.

20

Fig.2. Confocal image of autofluorescence of May-Grünwald-Giemsa-stained blood smear from a female coati. A: Hemoplasmas detected by autofluorescence of dye in blue. B: Differential interference contrast (DIC) image showing coccoid structures attached to erythrocytes, measuring approximately 0.4 µm in diameter (arrows). C: Composite image of the erythrocytes, detected by autofluorescence of dye in green, and hemoplasmas detected in blue. D: Composite image of the fluorescent and DIC images. Bar = 5 µm.

21

Fig.3. SEM images of blood collected from a female coati. A: Structures of about 0.3 µm in diameter are attached to erythrocytes (arrows). Bar = 3 µm. B: An organism in binary fission (arrowhead). Bar = 1 µm.

22

Fig. 4. Consensus tree of 8,251 trees from a Bayesian analysis of the 16S rRNA gene alignment of 44 hemoplasma sequences showing the relationship of coati mycoplasmas (in bold) and other hemoplasmas. BI posterior probabilities/ML bootstrap support values ≥70% are indicated at the nodes. Species names are followed by GenBank accession number, host and country of origin. The tree was rooted to Acholeplasma laidlawii (GenBank U14905). The scale bar represents the expected number of changes per site.