Correlation between presence of lactate dehydrogenase-elevating virus RNA and antigens in motor neurons and paralysis in infected C58 mice

195

Vhs Research, 6 ~~986~87)195-209 Elsevier

VRR 00297

Correlation between presence of lactate de~ydro~enas~-elevating virus RNA and antigens in motor neurons and paralysis in infected C58 mice Christopher H. Contag, Stephen P.K. Ghan, Stephen W, Wietgrefe and Peter G.W. Plagemann Department oj Microbiology, University ofMinnesota,Medical School, Minneapolis, MN 55455, U.S.A. (Accepted for publication 23 July 1986)

Summary

Lactate dehy~ogenase-elevat~g virus (LDV) induces poliomyelitis in i~unosuppressed CS8 mice resulting in fatal paralysis. We have synthesized and cloned cDNA complemental to the LDV genome, and used the cDNA clones as in situ hybridization probes for the detection of LDV RNA in tissue sections. Direct fluorescent antibody staining using IgG from chronically infected mice was used for the detection of LDV antigens. Using these methods, we have detected LDV RNA and antigens in anterior horn neurons of paralyzed mice. The appearance of LDV RNA and antigen positive motor neurons and their location in the spinal cord correlated with the development of paralytic symptoms. No positive neurons were detected in LDV-infected, susceptible mice without signs of paralysis, but some glial cells of the white and gray matter in the spinal cords of these mice were found to contain LDV RNA. These analyses broaden the host cell range of LDV to include neuronal and other cells in the CNS and support the hypothesis of LDV replication in neurons as the cause of poliomyelitis and paralysis. lactate dehydrogenase

elevating virus, in situ hybridization,

poliomyelitis, CS8 mice

The lactate dehydrogenase-elevating virus is an enveloped, positive-strand RNA virus that has been placed in an unnamed genus of the family of Togaviridae (B~nton-Da~ell and Plagemann, 1975; Brinton, 1980). Infections by LDV invaria~148-170~/$6/$03.~0~

1986

Elsevier

Science PublishersB.V.(Biomedical

Division)

196 bly result in a life-long, persistent infection of all strains of Mus musculus, which is associated with a continuous low level viremia (Rowson and Mahy, 1975; 1985). LDV has a very restricted host range both in vivo and within populations of primary macrophages in culture (Tong et al., 1977; Stueckemann et al., 1982). Although in most strains of mice infections by LDV are not associated with any clinical symptoms, under certain multifactorial conditions infections result in fatal paralytic poliomyelitis (Murphy et al., 1980; 1983; Martinez et al., 1980). Susceptibility to this motor neuron disease is linked to the Fv-1” allele (Murphy et al., 1980), which controls the expression of N-tropic retroviruses. In addition, several copies of endogenous N-tropic retroviral genomes must be present (Murphy et al., 1980). This genetic profile is found in AKR and 08 mice. Furthermore, susceptibility of C58 mice to LDV-triggered paralytic disease increases with age of the mice and is enhanced by immunosuppression (Murphy et al., 1980; 1983). The neurological disease is characterized by cytopathic changes in neurons in the anterior horn of the spinal cord and an influx of mononuclear cells (Murphy et al., 1980; Stroop and Brinton, 1983) but the mechanism by which LDV triggers these changes is not understood. LDV antigens have been reported to be detected in anterior horn neurons of the spinal cord of paralyzed C58 mice by indirect fluorescent antibody staining (Stroop and Brinton, 1985). However, unequivocal identification of the positive cells is not possible by this method and the question of staining, in the indirect method, of IgG and IgG carrying inflammatory cells, which generally surround degenerating neurons in paralyzed mice (see later), was not addressed. Also, the presence of LDV antigens in the CNS alone does not necessarily indicate a causal role of LDV in the induction of paralytic disease. To more clearly understand the pathogenesis of LDV in poliomyelitis and paralysis, we have cloned cDNA’s to the LDV genome and have used in situ hybridization with the cloned cDNA as well as direct fluorescent antibody staining with mouse anti-LDV IgG for the detection of LDV-infected cells. Cytochemical staining of sections of fixed tissues in combination with in situ hybridization allowed unequivocal identification of LDV RNA positive motor neurons in the anterior horns of paralyzed C58 mice. We have confirmed the presence of LDV antigens in the spinal cord and have further localized, in time and space, LDV antigen expression and the presence of LDV RNA in neurons. LDV positive neurons were confined to affected anterior horns of the spinal cord and their appearance correlated with the development of paralytic disease. In addition, LDV positive non-neuronal cells were detected in the white and grey matter of spinal cords of LDV-infected C58 mice, but their presence did not correlate with impairment of motor neuron function. The results indicate that LDV replication in anterior horn neurons plays a major role in the induction of paralytic disease. Materials and Methods Mice Outbred Swiss mice (about 4 weeks of age) were obtained from Biolabs, Inc. (St. Paul, MN) and C58/M mice from Dr. W.H. Murphy (University of Michigan). All

197 C5X mice were 6-8 months of age and immunosuppressed by intraperitoneal inoculation of 200 mg of cyclophosphamide/kg (Sigma, St. Louis, MO) 24-48 h prior to inoculation of about lo6 ID,, of LDV,/mouse. BALB/c mice were bred in the Department of Microbiology animal facility. LDV

The neurovirulent strain of LDV (LDV,) was obtained from Dr. W.H. Murphy (Murphy et al., 1980). LDV, was isolated in our laboratory (Brinton-Darnell and Plagemann, 1975). Con~ntrations of infectious LDV were estimated by end point dilution in mice (Plagemann et al., 1963). LDV stocks were prepared from pooled plasma of 50-100 20-22 h infected Swiss mice containing 109-lOi ID,,/ml and the virus was purified by isopycnic centrifugation in linear sucrose density gradients (Cafruny et al., 1982). Isolation of LD V RNA

Purified LDV (about 101’ ID,,) was incubated in a solution composed of 10 mM Tris-HCl (pH 7.4) and 1 mM EDTA containing 200 pg self-digested proteinase K/ml and 0.5% (w/v) sodium dodecyl sulfate at 37°C for 30 min. The mixture was then extracted twice with phenol/chloroform (1: 1) and once in chloroform alone. The RNA was precipitated with ethanol and stored under 70% ethanol at -70°C. Alternatively, LDV RNA was extracted from pu~fied LDV according to Cbirgwin et al. (1979) by extraction with gua~di~um isothiocyanate (Fluka, Switzerland) followed by centrifugation through a 5.7 M CsCl cushion in an SW50 rotor at 35000 rpm for 24 h. Generation of LDV cDNA probes

cDNA complementary to LDV RNA was synthesized according to standard protocols (Maniatis et al., 1982; Friedman et al., 1984) using random DNA primers. Random primers were generated from calf thymus DNA using DNase I (Worthington Biochemicals, Malvern, PA) digestion and ion-exchange chromatography on DEAE-cellulose (DE-52, Whatman). A mixture of RNA (about 2 pg) extracted from LDVr virions and 500 ng of random primers was heat denatured at 100°C for 2 min and slowly cooled to 42°C. Reverse transc~ption (11 U reverse transcriptase, Life Sciences, Inc., St. Petersburg, FL) was conducted at 42OC for 2 h in 50 mM Tris-HCl (pH 8.1), 50 mM KCl, 8 mM MgCl,, 10 mM dithiothreitot (DTT), 0.25 mM unlabeled deoxynucleotides (dATP, dCTP, dGTP and dTTP) and 0.7 mCi [a-32P]dCTP (3000 Ci/mmol)/ml. The reaction was terminated by addition of 20 mM EDTA, and the RNA hydrolyzed overnight in 0.3 M NaOH at 22OC. After neutralization, the cDNA was precipitated in ethanol and the second strand was synthesized with DNA polymerase KIenow fragment (2.5 U, Boehringer, Indianapolis, IN) and reverse transcriptase (11 U) in 32 mM Tris-HCl (pH 8.4), 107 mM KCl, 3 mM DTT and 0.1 mM of each of the four deoxynucleoside triphosphates. After purification by exclusion c~omato~aphy (Sephadex G-50, Pharmacia, Piscataway, NJ) and lyop~lization, single-str~ded regions were digested by incubation with S, nuclease (5 U; Boehringer Mannheim, F.R.G.) in 50 mM sodium acetate

198

(pH 4.6) 150 mM NaCi, 3 mM Zn(OAc), and 20 pg yeast RNA/ml for 30 min at 4°C (Sigma). The cDNA ends were filled in by incubation with Td polymerase (International Biotechnology, Inc., New Haven, CT) following the procedure recommended by the manufacturer. Blunt-ended cDNA was sized on a Sephacryl S-1000 column (Pharmacia, Piscataway, NJ) in 1 mM Tris-HCl/O.l mM EDTA. The fractions comprising the leading third of the peak were pooled and lyophilized. The final yield of cDNA from these fractions (about 600 ng) was blunt-end ligated into a Sma I-digested and phosphatased pUC 19 plasmid (Yanisch-Perron et al., 1985). Recombinants were selected by propagation on JMS3 E. coli(Yanisch-Perron et al., 1985). A plasmid contai~ng a 550 bp cDNA insert was selected by hyb~dization with LDV specific single-stranded [32P]cDNA. In addition, competent JM83 cells transformed with pUC 19-containing LDV cDNA inserts were propagated in batch liquid culture and the mixture of plasmids present in these cells was isolated and digested with endonucleases BamHI and EcoRI. Fragments of 1500-2500 bp (referred to as mixed probe) were size selected on 1.0% agarose gels, isolated by electrophoresis onto DEAE cellulose membranes (NA-45, Schleicher and Schuell; Fig. lA), and eluted according to the manufacturer’s protocol. The 550 bp cDNA referred to already was similarly isolated. The mixed probe and the 550-bp cDNA fragment were purified on ion-exchange columns (DE-52, Whatman) and used as in situ hyb~dization probes. LDV cDNA fragments were labeled with f1251]dCTP in standard nick translation reactions (Maniatis et al., 1982). Tissue preparations

Mice were anesthetized with ether and perfused with 60 ml of phosphate-buffered saline, pH 7.4 (PBS) while the mouse was encased in ice. The spinal cords were forced out of the spinal column by hydraulic pressure at the sacral end of the exposed decapitated vertebral columu. The spinal cord was divided into cervical, thoracic and lumbar regions, embedded in OCT compound (Tissue-Tek, Miles Scientific, Naperville, IL), ‘snap-frozen’ in liquid N, or between blocks of solid CO, and stored at - 70” C. Sections for imm~o~uorescence analysis were cut transversely on a cryostat at 6 pm thickness, picked up on glass slides, fixed by holding them for 30 min at room temperature and stored at - 70°C. Before use, the sections were immersed in acetone for IO min and air-dried. Frozen tissue sections of 8-10 pm for in situ hybridization were picked up on microscope slides that had been treated with Denhardt’s medium and acetylated. The slides were air dried, fixed in ethanol/acetic acid (3 : 1) and rinsed in absolute ethanol. Sections fixed in this manner were stored at 22°C for not more than two months. The preparation of aldehyde-fixed tissues was conducted as described by Brahic et al. (1984). Mice were perfused under ether anesthesia with 20 ml PBS, followed by 60 ml of ice-cold fixative containing 1% (w/v) parafo~aldehyde, 0.5% (v/v) glutaraldehyde, 0.1 M phosphate buffer (pH 6.0), 1.6% (w/v) r>-glucose, 0.~2~ (w/v) CaCl, and 1.0% (v/v) dimethylsulfo~de (PFG). During perfusion, animals

199 were encased in ice. Brain, spinal cord and spleen were removed and fixed further by immersion in PFG for 1 h followed by 70% ethanol for 1 h and 80% ethanol overnight. Tissues were processed for paraffin embedding using routine histological techniques. Tissues were embedded in Ameriffin (American Scientific Products, Minneapolis, MN). 6-pm thick sections were cut, floated on a 0.1% (w/v) solution of gelatin in water at 42OC, and picked up on microscope slides that had been treated with Denhardt’s medium and acetylated. The slides were dried at 37*C overnight and then stored at 4°C for not more than 2 weeks before hybridization. Hybridization to tissue sections

Sections of frozen tissue were pretreated for in situ hyb~d~ation as previously described by Haase et al. (1984). Briefly, sections were incubated for 20 min in 0.2 N HCl at 22”C, 30 min in 2 X SSC (0.15 M NaCl/O.O15 M sodium citrate) at 7O”C, digested at 37°C for 15 ruin with proteinase K (1 pg/ml of 20 mM Tris-HCl, pH 7.5/2 mM CaCl,) and incubated at 22°C for 10 min in 10 PM am-in tricarboxylic acid (ATA) in H,O. The sections were dehydrated in graded aqueous solutions of ethanol. Each section was overlayed with 5 ~1 of hybridization solution composed of 0.1 mg calf thymus DNA (50-1000 bp)/ml, 0.1 mg mouse liver RNA (50-500 bp)/ml, 50% (w/v) d eionized formamide, 5% (v/v) polyethylene glycol, 100 PM ATA, 0.6 M NaCI, 200 mM HEPES, 10 mM EDTA, 10 X Denhardt’s medium (Maniatis et al., 1982), 1 mg polyadenylic acid/ml and 0.4 pg ‘251-labeled LDVspecific cDNA/ml. The sections were then covered with a siliconized glass coverslip (18 mm diameter), this hyb~dization chamber was sealed with rubber cement around the perimeter of the coverslip and the hyb~dization carried out at 22OC for 3 days in the dark. After hyb~di~tion the slides were washed for 5 min in 1 x HWM (50% formamide, 10 mM HEPES, pH 7.2, 5 mM EDTA and 3 M NaCl), for l-2 min in 2 X SSC, for 1 h in 2 X SSC at 55°C and finally in 3 changes of 1 x HWM with constant stirring over a period of 3 days (Haase et al., 1984). The sections were rinsed twice in 2 x SSC and dehydrated in graded solutions of ethanol containing 0.3 M NH,OAc. Paraffin-embedded tissue sections were treated as described by Blum et al. (1984). Sections were incubated at 60°C for 3-4 h and then deparaffinized by soaking 3 times in xylene and twice in absolute ethanol (5 min each). The sections were rehydrated in 90% and 70% ethanol (5 min each), washed twice in PBS and treated with the following: 0.2 N HCI for 30 min at 22°C; 0.153 M t~ethanol~ne (pH 8.0) for 15 min at 22*C; 2 X SSC for 30 min at 70°C; 0.~5% (w/v) digitonin in 125 mM sucrose/60 mM KC1/3 mM HEPES @H 7.4) (26) for 5 min at 22°C; 5 pg proteinase K/ml of a solution of 20 mM Tris (pH 7.5) and 2 mM CaCl, for 15 min at 37*C; and finally 10 mM ATA for 15 min at 22°C. The sections were dehydrated by sequential exposure to 70%, 80% and 90% ethanol, air-dried and hybridized with cDNA as described already for frozen sections. After hybridization the slides were washed and autoradiographed as described by Haase et al. (1984). Slides were coated with nuclear track emulsion (NTB-2; Eastman Kodak, Rochester, NY) containing 0.3 M NH,OAc, air dried and exposed for 5-7 days. Slides were developed in Kodak D-19 developer, rinsed and fixed.

200 Sections of CNS tissues were stained with hematoxylin spleens were stained with bay-Gr~nw~d and Giemsa.

and eosin, and sections

of

Detection of viral antigen by direct immunojluorescence antibody staining A pool of serum from about 100 mice that had been infected with LDV, for 24 months was conjugated to fluorescein isothiocyanate according to the method described by Nairn (1969). The conjugate was then adsorbed with acetone-treated mouse liver and brain as well as murine derived tissue culture cells until non-specific staining of uninfected mouse tissue sections was completely abolished. Sections of frozen tissues were incubated with the FITC-labeled mouse anti-LDV for 40 min at 37°C in a humidified chamber, followed by 2 successive 10 min washes in PBS. After washing, the sections were mounted on coverslips with a drop of glycerol/ PBS (9 : 1) and examined with a fluorescence microscope.

Results

Synthesis and specificity of reagents for in situ hybridization and direct fluorescent antibody staining We have used both a 550 bp, cloned, LDV-specific cDNA and a cloned, mixed cDNA probe consisting of a mixture of fragments in the size range of 1500-2500 bp (Fig. lA), for in situ hybridization_ The cDNA was synthesized by random priming with small fragments of calf thymus DNA and cloned into a pUC19 plasmid. The recombinant plasmids were amp~fied as a mixed probe and the cDNA fragments were size selected. This mixed probe potentially covers the entire LDV genome and therefore enhances the sensitivity of detecting LDV RNA in tissue sections due to a complexity approaching that of the genome. The specificity of the mixed probe was assessed by hybridization to both northern blots and to 8-h LDV-infected and mock-infected primary macrophage cultures in situ. In northern blots of total RNA isolated from 8-h LDV-infected macrophage cultures, the cDNA probe specifically hybridized to an RNA with the molecular size of LDV RNA (about 12 kb, 48S, Brinton-Darnell and Plagemann, 1975) which was not present in total RNA extracted from uninfected macrophage cultures (Fig. 1B). Furthermore, the cDNA probe hybridized specifically to the subpopulation of macrophages in primary cultures that are LDV permissive and generally make up between 5 and 15% of the total cells in these cultures (Tong et al., 1977). A similar proportion of cultured macrophages was specifically stained by direct fluorescent antibody staining (data not shown). The macrophages in the spleen are probably a major site of LDV replication, at least during the acute phase of infection (Plagemann and Swim, 1966; Porter et al., 1969; Kowalchyk and Plagemann, 1985). Thus, to assess the ability of our methods to detect LDV RNA as well as antigens in tissues, we have examined sections of frozen spleens from l-day infected BALB/c mice for LDV RNA and antigens by in situ hybridization with the 550 bp cDNA probe and by direct fluorescent antibody staining. Both procedures revealed numerous foci of positive cells in frozen sections

201

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Fig. 1. Agarose gel electrophoresis of LDV-specific inserts of 1500-2500 bp (A) and hybridization of this cDNA probe to RNA from l-day LDV-infected and uninfected primary macrophage cultures (B). (A) Plasmids were isolated from transformed JM83 cells which had been propagated in batch liquid culture. The inserts were recovered by digestion with BamHI and EcoRI, size selected by electrophoresis on 1% agarose gels and purified (see Materials and Methods). The mixed cDNA probe was electrophoresed on a 1% agarose gel for confirmation of size (lane 1). The molecular weight markers (lane 2) consisted of a mixture of HindIII-digested phage h DNA and HaeII-digested replicative form of #X174 (DRIgest, Pharmacia). (B) Total RNA (1 pg) from 8-h LDV-infected (lane 1) and uninfected (lane 2) macrophage cultures was electrophoresed on formaldehyde agarose gels and transferred to nitrocellulose. 32P-labeled cDNA was hybridized under stringent conditions and the blot was autoradiographed.

(Fig. 2A and C). The pretreatments required for the detection of LDV antigens in tissue sections have precluded the simultaneous detection of antigens and genomes of LDV. We are currently working out a protocol which is suitable for the detection of both the intracellular LDV antigens and RNA. Both the LDV RNA- and LDV antigen-positive cells were located most often in the red pulp of the spleen, the region most heavily populated by macrophages. No LDV-positive cells were observed in sections of spleens from uninfected mice (Fig. 2B and D). Incubation of the spleen sections with RNase A and Tl before in situ hybridization reduced the signal to background (data not shown). The results demonstrate the reliability of both methods for detecting cells containing LDV RNA and antigens in tissue sections. There was good agreement between the distribution of cells positive for LDV RNA and positive for LDV antigens in the spleen sections (Fig. 2). The positive cells could not be identified as

Fig. 2. In situ hybridization and direct fluorescent antibody staining of sections of spleen from 24-h LDV,-infected (A and C) and uninfected (B and D) BALB/c mice. Spleen sections (8 pm thickness) were hybridized in situ with a LDV-specific [I*’ I]cDNA fragment of 550 bp in length (specific activity approximately 5~10~ cpm/pg), exposed for 5 days, developed and stained with May-Grtinwald and Giemsa stains. (A and B, magnification: 800 x .) Other 6 pm sections were examined by direct fluorescent antibody staining as described under Materials and Methods. (C and D, magnification: 620~ .) Arrows to autoradiographic grain clusters in (A) denote infected cells,

203 to cell type, but they probably are productively-infected macrophages since the spleen is known to contain numerous macrophages that support LDV replication in vitro and macrophages are the only type of LDV-permissive cell that can be cultured from the spleen (Plagemann and Swim, 1966; Kowalchyk and Plagemann, 1985). In earlier experiments (Kowalchyk and Plagemann, 1985; Cafruny et al., 1986a), we measured the distribution of LDV antigens in infected macrophage cultures by indirect fluorescent antibody staining using mouse anti-LDV IgG and FITC rabbit anti-mouse IgG. We found this method far less suitable for detecting LDV antigenpositive cells in spleen sections than the direct method, because of considerable background staining of the sections, presumably due to the presence of mouse IgG and IgG containing cells in tissues. This obviously complicates indirect fluorescent antibody analysis of spinal cords containing inflammatory infiltrates. Presence of LDV RNA and antigens in anterior horn neurons of paralytic C58 mice 6- to S-month old C58 mice were immunosuppressed by injection of cyclophosphamide and then infected with lo6 ID,, of LDVJmouse. Under these conditions greater than 80% of the mice develop paralytic symptoms lo-22 days p.i. (Murphy et al., 1980; Martinez et al., 1980; Cafruny et al., 1986b). In initial experiments, mice were sacrificed at the height of paralytic disease. Their spinal cords and brains were quick frozen, sectioned and the sections were examined by in situ hybridization or by direct fluorescent antibody staining. In situ hybridization of the mixed cDNA probe to sections of frozen spinal cord revealed numerous foci of autoradiographic grains in transverse sections of the lumbar region (Fig. 3A). No foci of autoradiographic grains were detected in RNase A and Tl-treated sections of spinal cords from paralyzed mice or in sections of spinal cords from uninfected mice in numerous experiments (data not shown). Similarly, we detected numerous LDV antigen-positive cells by direct fluorescent antibody staining in these sections of frozen spinal cords of paralyzed C58 mice (Fig. 3B), whereas none were detected in sections of spinal cords from uninfected mice (data not shown). The LDV antigen-positive cells were confined to the lumbar region and their frequency was comparable to the distribution of LDV RNA-containing cells detected by in situ hybridization. The morphology of cells in the frozen sections of spinal cords was not sufficiently preserved to identify the cell type of the LDV-positive cells. In order to enhance the preservation of cellular morphology, we fixed the spinal cords in a solution containing paraformaldehyde and glutaraldehyde (PFG). The fixed tissues were embedded in Ameriffin, cut into sections of 6 pm thickness, and the sections were hybridized with the mixed cDNA probe. Cellular morphology was sufficiently preserved in these experiments to allow definitive identification of LDV RNA-positive motor neurons in the anterior horn of the lumbar region (Fig. 4A, B). Most LDV RNA-positive neurons were surrounded by other types of cells (Fig. 4A, B), which we suspect represent inflammatory cells. Many neurons without LDV RNA were also present in the lumbar region and these were not associated with inflammatory cells (Fig. 4A, C). In addition, non-neuronal cells in various regions of

204

Fig. 3. In situ hybridization (A) and direct fluorescent antibody staining (B) of sections of frozen spinal cord from a paralyzed mouse. The spinal cord was removed at the height of paralysis (15 days pi.), quick-frozen and sections of 8 pm thickness were cut. (A) Sections were hybridized in situ with an LDV-specific ‘251-labeled mixed cDNA probe (specific activity approximately 3 X lo9 cpm/pg), exposed for 7 days, developed and stained with hematoxylin and eosin. The photograph shows multiple foci of autoradiographic grains in one anterior horn region of the spinal cord. The posterior region as well as the other anterior horn contained few, if any, foci. Magnification: 600X. (B) Sections were examined by direct fluorescent antibody staining as described under Materials and Methods. The photograph shows LDV antigen-positive cells in the lumbar region of the spinal cord. Magnification: 440 x

the spinal cord were found to contain LDV RNA (Fig. 4C), but no positive cells were detected in sections of the brain and brain stem (data not shown). Sections of PFG fixed tissues or macrophage cultures were found to be unsuitable for direct fluorescent antibody staining as also reported for indirect fluorescent antibody staining by Stroop and Brinton (1985).

Fig. 4. Detection of LDV-infected cells in: the white and gray matte+ of spinal cords of LDV,-infected C58 mice by in situ hybridization. Longitudinal sections (6 pm thickness) of’ PFG-fixed paraffin embedded spinal cords from C58 mice were hybridized in situ with a LDV-specific t251-labeled mixed cDNA probe (specific activity approximately 2X109 cpm/pg), exposed for 7 days, developed and stained with hematoxylin and eosin. (A-C) Sections of spinal cord lumbar region taken from a mouse 15 days p.i. at the height of paralysis. (A) Anterior horn neuron with autoradiographic grains and an anterior horn neuron without grains among non-neuronal cells in the lumbar region, Magnification: 400 X . (B) Higher magnification of another LDV RNA positive neuron. Magnification: 625 x (C) Glial cell of the anterior horn with autoradiographic grains and a motor neuron without grains. Magnification: 325 X (D) Positive glial cells in the white matter of the lumbar region of the spinal cord of a C58 mouse that showed slight hind leg weakness, but no paralysis at 7 weeks pi. Magnification: 325 x

205

206 Temporal and spatial distribution of LDV-infected cells in the CNS In transverse sections of spinal cords from C58 mice with paralysis in a single hind leg, many foci of autoradiographic grains were detected in one of the anterior horns while few, scattered foci were present in the opposite anterior horn and other regions of the frozen spinal cord sections (data not shown). This correlation between the distribution of LDV-infected neurons and asymmetric paralysis was observed in three mice. Moreover, in longitudinal sections of PFG-fixed paraffin-embedded spinal cords from mice demonstrating only hind leg paralysis, motor neurons containing LDV RNA were always located in the lumbar region, and no LDV positive neurons were found in any other region of the spinal cord from three paralyzed C58 mice although numerous sections were examined. We also did not detect any neurons containing LDV RNA in any region of longitudinal sections of spinal cords of 24-h and 5-day LDV-infected, cyclophosphamide-treated C58 mice or in cyclophosphamide-treated C58 mice that had not developed paralytic symptoms by 7 weeks p.i. There were, however, other types of cells present that contained LDV RNA (Fig. 4D). These cells were observed in both the white and gray matter of the spinal cord.

Discussion The mixed LDV-specific probe we have prepared is a double-stranded, cloned version of the probes generated by incorporation of radiolabeled precursors into single-stranded cDNA during reverse transcription that are generally used for in situ hybridization (Haase et al., 1984). Although we may have sacrificed hybridization efficiency by using a double-stranded probe, we have generated an amplifiable library of probes by cloning the cDNA into a high copy number bacterial plasmid. Any decrease in the efficiency of hybridization can be compensated for by the addition of high molecular weight polymers, such as polyethylene glycol (Wahl et al., 1979; Wetmur, 1975) and denaturing agents (e.g., formamide) to the hybridization solution (McConaughy et al., 1969). We have, therefore, created an inexhaustible supply of cDNA probes without suffering serious losses in hybridization efficiency or specificity. In situ hybridization, using the mixed probe, produces signals of much greater intensity than small single fragments (cf. Fig. 1A and Fig. 3). We are, at present, sorting out the cloned fragments to determine the exact complexity represented by the mixed probe and to discern the regions of the genome that are represented in this library. The cloned cDNA fragments also provide a set approach to rapidly attain of clones, which can be sequenced in a ‘shotgun’ information on the genomic sequence of LDV. The in situ hybridization, using the mixed probe, allowed unequivocal identification of LDV-infected anterior horn neurons in paralyzed C58 mice. Furthermore, the spatial as well as temporal correlations between the development of paralytic symptoms in LDV-infected C58 mice and the presence of LDV-RNA in anterior horn neurons suggest a causal relationship. However, it is still unclear whether motor neuron dysfunction results simply from the cytocidal replication of LDV in

207 these cells. Although both LDV RNA and antigens are found in the spinal cord of paralytic mice it has not been established yet that LDV RNA and protein are always present in the same neurons nor that intact virions are produced. Electron microscopic studies have failed to detect mature virions in spinal cord motor neurons of paralytic C58 mice (Stroop and Brinton, 1985). It is possible that non-productive infections cause dysfunction of motor neurons. Motor neuron destruction could also involve a host immune response triggered by the replication of LDV in these cells, as suggested by reports of neurophagia (Stroop and Brinton, 1983) and the frequent presence of inflammatory cells surrounding LDV RNA-positive neurons (Fig. 4A, B). Our results indicate that LDV RNA-positive neurons appear in the spinal cord sometime after 5 days p.i., but a larger number of animals needs to be analyzed to more clearly delineate the time course of spread of LDV to the CNS. In addition, the identity of the various non-neuronal cells in the spinal cord that become infected needs to be ascertained. Since LDV RNA-positive non-neuronal cells, but not neurons, were detected in mice without paralytic symptoms, it seems likely that it is not simply the spread of LDV to the CNS that determines whether or not motor neurons become infected. At present the mechanism of spread of LDV to the CNS, the types of non-neuronal cells that become infected and the factors that may affect the infection of neurons are uncertain and difficult to ascertain. It is possible that infected macrophages may transport LDV to the CNS, as has been suggested for visna virus (Peluso et al., 1985), although other mechanisms may be involved. The LDV RNA-positive, LDV antigen-positive non-neuronal cells in the spinal cord may be glial cells that are LDV permissive. In situ hybridization combined with immunocytochemical examination for cell specific surface antigens may yield some information on the identity of the LDV RNA-positive, non-neuronal cells and the mechanism of spread of LDV into the CNS. Indirectly our results suggest the presence of surface components on neurons of C58 mice that function as receptors for LDV infection, but whether or not these are the same as the LDV receptors on permissive macrophages (Kowalchyk and Plagemann, 1985) is unknown. A related question is whether the unique susceptibility of C58 and AKR mice to LDV-triggered paralytic disease is due to the presence of specific LDV receptors on motor neurons of these mice, but not on those of non-susceptible mouse strains. Of interest in this respect is the finding that various isolates of LDV differ in neurovirulence and that repeated passage of an LDV strain through C58 mice enhances its neurovirulence for these mice (Murphy et al., 1980, 1983). It is possible that passage through C58 mice may select LDV variants with increased affinity for surface receptors on neurons of these mice. The question of cell specificity is complicated by the fact that the restriction of LDV-triggered paralytic disease to C58 and AKR correlates with the presence of several copies of an endogenous ecotropic retrovirus and permissiveness to retrovirus expression (Murphy et al., 1980, 1983). What, if any, function endogenous retrovirus expression may have in rendering these mice sensitive to LDV-triggered paralytic disease is an unsolved question. Equally unclear is the mechanism by which immunosuppression enhances the sensitivity of C58 mice to the disease and how certain lymphocytes

208 from young C.58 mice protect old C58 mice from developing symptoms (Mushy al., 1980, 1983).

et

Acknowledgements

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