Viremia has been detected in many herpesvirus-infected species, including horses,1–4 humans,5, 6 guinea pigs,7 pigs,8 sheep,9 and cows,10 and is important in the pathogenesis of herpetic disease in these species. In contrast, the role of viremia in the pathogenesis of herpetic disease in cats is not well understood. Although the virus is readily isolated from conjunctival, nasal, and oropharyngeal samples of affected cats,11–15 there are infrequent reports of isolation of FHV-1 from blood12, 16 or internal organs11,17,18 of cats infected via natural routes. For these reasons, viremia has been reputed to play a minor role in dissemination and later reactivation of FHV-1. Rather, FHV-1 is believed to reach the trigeminal ganglia by ascending sensory nerve axons.13, 19 Subsequent reactivation of latent virus within the ganglia is thought to be followed by retrograde delivery of viral particles along the same sensory nerves to peripheral epithelial sites where the virus can be readily isolated. This so-called round-trip theory has led to the assumption that viremia is unimportant in the pathogenesis of FHV-1 in cats. In other species infected with herpesviruses and in cats infected with viruses other than FHV-1, viremia allows dissemination of the virus to all organs and can cause abortion, encephalitis, disseminated intravascular coagulation, and generalized organ failure.8,20–22 Therefore, further understanding of the role of viremia in the pathogenesis of feline herpetic disease is important.
The ability to detect viremia has been improved by the wide availability and high sensitivity of PCR assays. Although PCR assays have been used to detect FHV-1, typically only samples collected from peripheral anatomic sites such as skin, conjunctiva, cornea, and the oral cavity of cats with clinical evidence of primary or recrudescent herpetic disease have been tested.14,15,ref23–29 Less commonly, FHV-1 DNA has been detected in the blood of cats with clinical signs of herpetic disease30, 31 as well as apparently clinically normal cats.32 However, this has not been a consistent finding.23 The lack of studies describing detection of FHV-1 in serial blood samples from experimentally infected cats, along with the lack of studies in which viable virus is isolated from feline blood, makes it difficult to accurately define the role of viremia in recrudescent FHV-1 disease. Therefore, the purpose of the study reported here was to determine the presence of FHV-1 in the blood of cats undergoing primary experimentally induced herpetic disease and in cats undergoing presumed FHV-1 recrudescence following natural infection, most of which were shedding FHV-1 in their conjunctival fornix.
Materials and Methods
Experimental primary FHV-1 infection—Four male and 2 female unvaccinated SPF domestic shorthair cats approximately 6 months of age and without known history of ocular or systemic illness were included in this study. All animals were maintained and handled in accordance with the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research, and all experimental procedures were approved by the University of California, Davis, Institutional Animal Care and Use Committee. Prior to inclusion in the present study, all cats were determined not to have detectable FeLV antigena or antibodies against FIVa or FHV-1b in blood samples. Following a 24-hour period of acclimatization, all cats underwent complete general physical and ophthalmic examinations. General physical examination included assessment of rectal body temperature and pulse and respiratory rates, along with thoracic auscultation and clinical assessment of hydration, mucous membrane color, and behavior. Ophthalmic examination included slit-lamp biomicroscopy and assessment of direct and consensual pupillary light reflexes, palpebral reflex, and menace response. All cats were also assessed via CBC, serum biochemical analyses, and urinalysis. Examination findings and test results were within reference ranges for all cats. Following collection of baseline data, all cats were inoculated topically in both nares and both conjunctival fornices, with a total of approximately 3.2 × 107 pfu of strain 727 (passage 8) FHV-1. This virus is a plaque-purified field isolate that has been verified as FHV-1 by use of results of immunofluorescence with FHV-1–specific antiserum13 and PCR assay15 and verified as not contaminated by Mycoplasma spp,b Chlamydophila felis,c or feline calicivirus,c which may have confounded the clinical disease induced in the study. No cats received any topical (ophthalmic) or systemic treatment during the study. In the final week of the study, FHV-1 serologic examinations, CBC, serum biochemical analyses, and urinalysis were repeated.
One day prior to inoculation and once daily following inoculation for 21 days, clinical signs of disease attributable to FHV-1 infection were scored according to a published scale.26 Severity of conjunctivitis was assigned a score of 0 (none) through 3 (moderate to severe). Severity of blepharospasm was assigned a score of 0 (none) through 4 (eye completely closed). Severity of ocular discharge was assigned a score of 0 (none) through 3 (marked mucopurulent discharge). Sneezing was assigned a score of 0 (absent) or 1 (present). Severity of nasal discharge was assigned a score of 0 (none) through 3 (marked mucopurulent discharge). Clinical score for each eye was defined as the sum of all ocular (conjunctivitis, blepharospasm, and ocular discharge) and nonocular (sneezing and nasal discharge) scores. Total clinical disease scores were defined as the sum of scores for all scored categories for each cat, with the minimum and maximum possible total clinical score being 0 and 14, respectively. As part of another unpublished study, cats also underwent biopsy of the ventral conjunctival fornix immediately prior to FHV-1 inoculation and on PIDs 7, 14, and 21. Periodically, from prior to inoculation until PID 21, blood was collected by jugular venipuncture from a variable number of cats. The timing and number of cats sampled were as follows: prior to inoculation (n = 6), PID 1 (3), PID 2 (3), PID 3 (6), PID 4 (3), PID 6 (5), PID 8 (2), PID 9 (1), PID 12 (5), PID 15 (6), PID 19 (1), PID 20 (1), and PID 21 (3). Sampling intervals were dictated by use of the protocol of another unpublished study in which these cats were involved. In all cats, blood was collected by jugular venipuncture into a commercial blood collection tube containing sodium EDTAd and stored at 4°C for subsequent analysis via PCR assay. For DNA extraction, blood samples were gently inverted prior to sterile transfer of a 100-ML aliquot to a 1.5-mL microcentrifuge tube. The DNA from these samples was then purified by use of a commercially available kit,e and 300 ng of the extracted DNA was subjected to a nested, traditional end point PCR assay targeting a 224-bp region of the FHV-1 thymidine kinase gene, as described.27 This PCR assay reliably detects as few as 0.6 genomic copies of viral DNA and is highly sensitive, compared with other PCR assays for detection of this virus.33 Negative controls (PBS solution) were always included during DNA extraction. Negative (water only) and positive (purified FHV-1 DNA) controls were always included during thermocycling. As an additional positive control, prior to infection, a blood sample from 1 cat was collected into a tube containing sodium EDTA, and 100-ML aliquots were spiked with 8 × 103 pfu, 8 × 102 pfu, or 80 pfu of strain 727 FHV-1. These were then processed as for test samples.
Presumed herpetic recrudescence in naturally infected cats—Thirty-four adult cats from the sick bay of a local animal shelter, some of which had clinical signs suggestive of herpetic disease, were selected for inclusion in this study. All cats were assigned a total disease score for clinical signs of potential herpetic disease by a single observer (HDW) using the same published scoring system as in the initial part of the present study.26 Following clinical scoring, 0.5% proparacaine was applied to each eye and a separate swabf was rolled vigorously in the ventral conjunctival fornix of each eye. Each swab was then placed in a dry 1.5-mL microcentrifuge tube and stored on ice for 1 to 3 hours before being frozen at −20°C until DNA extraction.
Blood from each cat was collected by jugular venipuncture into a commercial CPTg and placed immediately on ice for 1 to 3 hours until separation of peripheral blood mononuclear cells was performed. After allowing the blood to equilibrate to room temperature (approx 22°C) for 10 to 20 minutes, peripheral blood mononuclear cells were collected from each CPT according to the manufacturer's instructions and resuspended in 1 mL of PBS. The mononuclear cell concentration in each sample was determined with a hemacytometer.h A portion of the suspension was immediately inoculated onto confluent CRFK cell monolayers. For 19 of the 34 samples, 250 ML of the mononuclear cell suspension was inoculated onto CRFK cell monolayers in a 6-well tissue culture plate and incubated in Dulbecco modified Eagle medium at 37°C with 5% CO2 for 6 days. Two replicates were performed for each sample. The remaining 15 samples were cultured by use of an alternate technique to enhance the chance of isolating virus that may have been latent within monocytes.2, 34 For those samples, 250 ML of the mononuclear cell suspension was inoculated onto a CRFK cell monolayer in a 25-cm2 flask. Another 250 ML of the cellular suspension was inoculated onto CRFK cell monolayers in Dulbecco modified Eagle medium supplemented with hydrocortisonei to a final concentration of 1 μg/mL. Both flasks were incubated at 37°C with 5% CO2 for at least 5 days and then passaged. Passaging was carried out by submitting the cells to 3 rapid freeze-thaw cycles, then pulse vortexing for 10 seconds, and inoculating 1 mL of the resulting lysate onto fresh confluent CRFK cell monolayers with or without hydrocortisone. All samples were passaged 4 times.
Positive and negative control samples were also submitted to both VI techniques. Negative controls consisted of 250 ML of sterile PBS solution. Positive controls consisted of 250 ML of medium containing 30 to 3,000 pfu of strain 727 (passage 9) FHV-1. As an additional positive control, 4 blood samples from 2 apparently healthy cats were collected into CPTs and placed on ice for 1 to 3 hours. Then 0 pfu, 3 × 102 pfu, 3 × 105 pfu, or 3 × 107 pfu of strain 727 (passage 9) FHV-1 was added to each sample. Mononuclear cells were harvested from these spiked blood samples and inoculated onto a confluent CRFK cell monolayer and incubated for 5 days in hydrocortisone-free media. All cell cultures inoculated with control and test samples were examined every other day for the appearance of cytopathic effects.
A commercial DNA extraction kitj was used to extract DNA from the conjunctival swab specimens and from 100 ML of those mononuclear cell suspensions that contained > 2 × 103 cells/mL. The presence of FHV-1 DNA within each sample was then determined by use of the same PCR assay described for the experimentally infected cats.27 Negative controls (PBS solution) were always included during DNA extraction. Negative (water only) and positive (purified FHV-1 DNA) controls were always included during thermocycling. As an additional positive control, blood samples from 2 apparently healthy cats were collected into 4 CPTs and placed on ice for approximately 30 minutes. Then 1 CPT each was spiked with 0 pfu, 3 × 102 pfu, 3 × 105 pfu, or 3 × 107 pfu of strain 727 (passage 9) FHV-1. Mononuclear cell suspensions from these samples were processed as for test subjects. If FHV-1 DNA was detected in only 1 of the 2 conjunctival swabs collected from each cat, only the swab in which DNA was detected was considered for statistical analysis. A Mann-Whitney rank sum test was used to assess the relationship between clinical scores and presence of FHV-1 DNA in conjunctival swabs. Significance was set at P b 0.05.
Results
Experimental primary FHV-1 infection—Prior to inoculation, all 6 SPF cats were clinically normal (total disease score, 0) and had no detectable serum antibodies against FHV-1. By PID 21, all cats developed dendritic corneal ulcers, had other clinical signs typical of FHV-1 infection, and seroconverted with respect to FHV-1. Median total disease score increased to a peak of 9.5 on PID 7 and then gradually declined. By use of the PCR assay, FHV-1 DNA was detected in blood of all 6 cats on at least 1 occasion. Considering all cats together, FHV-1 DNA was detected in blood on 10 occasions from PID 2 through 15. Nine of these 10 viremic episodes were detected from PID 2 through 9; the final viremic episode was on PID 15. Herpetic DNA was detected in the blood on only 1 day in 4 cats, on 2 days in 1 cat, and on 4 days in 1 cat. By use of PCR assay, FHV-1 was detected in all spiked blood samples, but in no negative control samples. All cats developed leukocytosis, as expected for primary FHV-1 infection. However, there were no changes detected via serum biochemical analyses or urinalysis that suggested renal, hepatic, or other specific organ involvement.
Presumed herpetic recrudescence in naturally infected cats—Clinical scores for the 34 adult cats from the shelter's sick bay ranged from 0 to 11 (median, 3). Feline herpesvirus type 1 DNA was detected in conjunctival swabs collected from 27 of the 34 (79.4%) cats. Of those, FHV-1 was detected bilaterally in 14 of 27 (52%) cats. Clinical signs consistent with herpetic infection were seen in 25 (92.6%) cats in which FHV-1 DNA was detected. Six cats did not have any clinical signs of herpetic disease (total disease score, 0). Feline herpesvirus type 1 DNA was detected in conjunctival swabs from 2 of these 6 cats. A significant difference in median clinical score was not detected (P = 0.509) between cats from which FHV-1 DNA was (median clinical score, 4) or was not (median clinical score, 0) detected in the conjunctival fornix.
Blood samples were collected from all 34 adult shelter cats; however, mononuclear cells could not be isolated from 3 samples. Median mononuclear cell concentration in the remaining 31 samples was 6 × 105 cells/mL (range, 2 × 103 cells/mL to 3 × 106 cells/mL). Cytopathic effects typical of FHV-1 were not induced in CRFK cell monolayers inoculated with any mononuclear cell samples, regardless of the number of times lysates were passaged or whether hydrocortisone was present in the culture medium. Similarly, FHV-1 DNA was not detected by use of PCR assay in any of the 31 mononuclear cell isolates tested. Cytopathic effects characteristic of FHV-1 were seen in CRFK cell monolayers within 3 days following inoculation with positive control samples containing native virus or mononuclear cell isolates spiked with 3 × 105 pfu and 3 × 107 pfu of FHV-1 but not with 3 × 102 pfu. Negative control samples were always negative by VI. By use of PCR assay, FHV-1 was detected in all spiked mononuclear cell isolates, but not in negative control samples.
Discussion
In the present study, it was possible to detect FHV-1 DNA sporadically in the blood of cats for approximately 2 weeks following experimental primary infection. These results support those of a previous study16 in which FHV-1 was isolated for a limited period from the blood of some cats undergoing experimental primary herpetic disease. However, there are a number of differences in methodology between the 2 studies that likely affect the comparison. In the previous study,16 viable virus was isolated on PID 8 from separated mononuclear cells in 50% of cats, whereas in the present study, viral DNA was detected between PID 2 and 15 in whole blood of every cat tested. The difference in detection rates between the 2 studies likely reflects, at least in part, the greater sensitivity of PCR assay versus VI.4,24,27,33 However, it is possible that viral viability also contributed to the difference. Because PCR assay detects only DNA, nonviable virus or potentially even viral fragments may have been detected in blood from cats in the present study and may explain the higher detection rate and longer detection period, compared with those in the previous study.16 Unfortunately, viral viability in blood collected from the experimentally infected cats was not determined because VI was not performed during that phase of the study. Additionally, in that phase of the present study, PCR assay was performed on whole blood, whereas in the previous study,16 virus was isolated from mononuclear cells only. Because FHV-1 is a cell-associated virus, it would be expected to be found in blood within mononuclear cells. However, cell lysis as a result of viral infection or during sample processing may have liberated virus from mononuclear cells and enhanced viral detection from whole blood in that phase of the study. Finally, because cats in the present study were participating in a separate unpublished study requiring weekly conjunctival biopsies, the present study's methodology did not truly reflect natural primary infection. It is conceivable that breaches in the conjunctival epithelial barrier induced by biopsy enhanced viral access to the bloodstream. However, natural primary FHV-1 infection in cats can cause ulcerative conjunctivitis, sometimes with hemorrhagic ocular discharge, suggesting that some degree of exposure of viral particles to blood within conjunctival capillaries occurs.
In light of the present results, it is interesting to consider what role viremia might play in cats undergoing primary FHV-1 infection. Certainly, this means of viral dissemination would expose many more tissues to potential viral infection than is expected via the accepted axonal means of spread. This in turn might be associated with abortion, disseminated disease such as pneumonia or encephalitis, or congenital (transplacental or parturient) infection. However, clinically evident involvement of tissues outside the respiratory tract and eyes is not reported in cats experimentally12, 35 or naturally36 infected with FHV-1 via a mucosal route, and cats in the present study had no clinical or clinicopathologic evidence of internal organ involvement during the 21 days following inoculation. Perhaps the strongest data suggesting that internal organ involvement is unlikely despite viremia are available from studies35, 37 in which neonatal, weanling, and adult cats were inoculated IV with FHV-1. Virus was subsequently detected in the blood, liver, spleen, lungs, bone, placenta, and uterus of many of these cats. However, only some cats inoculated IV developed clinical signs typical of natural FHV-1 infection, and these were mild and appeared to be age dependent. Weanling and adult cats failed to develop severe clinical signs or histologic lesions typical of herpetic upper respiratory tract disease,35, 37 and weanlings inoculated by the same route developed relatively mild signs, compared with kittens inoculated mucosally at that age.37 Taken together, data in the present study and from previous reports suggest that viremia does occur during primary infection via the mucosal route; however, visceral or neurologic involvement seems unlikely. This may be a manifestation of viral tropism.
In contrast to the data generated for cats undergoing experimental primary infection, it was not possible to detect FHV-1 DNA or to isolate live virus in blood samples from any of the adult shelter cats despite the fact that most of the cats had detectable FHV-1 DNA in their conjunctival fornices at the time of blood collection. The inability to detect FHV-1 DNA in the blood may be interpreted in a number of ways. It is possible that the collection method prohibited detection of virus. In this part of the study, we attempted to detect virus in blood mononuclear cells isolated by use of a commercially available CPT. This was done for a number of reasons. Data from the cats with primary infection revealed that virus was detected relatively infrequently and for a short period during primary infection. It was hypothesized that, as for peripheral viral shedding, the number of viral particles present intravascularly in recrudescent disease was likely to be lower than that present during primary disease. Also, it was hypothesized that, as in epithelial and neural cells, FHV-1 would likely be found in the nucleus. Therefore, viral detection was performed by isolating peripheral blood mononuclear cells. Moreover, because blood is not an adequate substrate to use for VI, isolation of peripheral blood mononuclear cells allowed VI and PCR assay on the same substrate, thereby maintaining consistency of data from these 2 techniques within the shelter cat population. To the authors' knowledge, a CPT has not been reported for this use in cats, and it is possible that the tubes contained inhibitors of virus or PCR that prevented VI or DNA detection. However, this method has been used to successfully isolate equine herpesvirus and to harvest herpesviral DNA from horses.2 Finally, isolation of FHV-1 and detection of FHV-1 DNA from spiked mononuclear cell samples in the present study suggested that the tubes do not completely inactivate FHV-1 or fully inhibit the PCR reaction. It is also possible that the cats tested in the second part of the study were not latently infected with FHV-1. The assumption that most cats in the shelter population were likely FHV-1 carriers was based on knowledge that at least 97% of adult cats are seropositive for FHV-138 and that at least 80% of FHV-1-infected cats become latently infected.11 Additionally, FHV-1 infection is common in shelters. Other studies of shelter populations revealed that FHV-1 DNA can be detected in 23% to 52% of cats39, 40 and that at least half of those cats shed virus within 7 days of entering a shelter.40 In cats with signs of upper respiratory tract disease, FHV-1 DNA detection rates can be as high as 92%.39 This has led to the conclusion that adult cats typically are latently infected and undergo viral reactivation upon entering shelters.10 Data from the present study support this hypothesis because FHV-1 DNA was detected in the conjunctival fornices of approximately 80% of the shelter cats tested, and most had clinical signs consistent with herpetic infection. On the basis of all of these data, it seems reasonable to assume that most cats tested in that part of the present study were latently infected with FHV-1.
It is possible that the inability to detect virus or viral DNA in blood samples from cats in the second part of the present study resulted from only 1 sample being collected from each cat. Considering the fairly limited period during which FHV-1 was detected in the blood of cats undergoing primary infection in this and a previous study,16 it appears that viremia occurs during a limited period only. It is possible that viremia during recrudescent disease is similarly limited and that we failed to detect virus because of sampling artifact. However, this possibility seems unlikely considering the number of cats sampled at random time points after their admission to the shelter sick bay and the range of clinical disease severity observed. Nevertheless, longitudinal studies in cats undergoing recrudescent disease are necessary to further investigate this possibility.
Our data suggest that viremia does occur in cats undergoing primary herpetic infection but not in cats undergoing recrudescent herpetic disease and that this likely reflects a true difference in the biology of this virus during those 2 phases of this infectious disease. This is in stark contrast to data from 3 studies30–32 in which herpetic viremia was examined by use of PCR assay in a total of 41 cats. None of those reports permits readers to distinguish cats undergoing primary versus recrudescent disease. However, all cats were r 6 months of age, and all but 1 cat30 were healthy32 or affected by ocular disease in the absence of respiratory signs.32 Therefore, it is unlikely that they were undergoing primary exposure to FHV-1. Of those 41 cats, 18 were healthy,32 of which 5 were viremic. Of the 5 viremic cats, 2 also were shedding FHV-1 in at least 1 conjunctival fornix and 1 also had DNA of Chlamydophila felis or Mycoplasma spp detected in at least 1 conjunctival fornix. Of the 23 cats with keratitis, conjunctivitis, or keratoconjunctivitis, 14 were viremic and 7 of those were shedding FHV-1 at the ocular surface.30, 31 No attempt was made to isolate viable virus from the blood of any cats in those 3 studies.30–32 The reason for the differences in detection rates of viremia between those studies and the present study is not known but may represent differences in handling, DNA extraction, or thermocycling techniques, as these are not described for the previous studies.
Taken together, results of the present study suggest that a brief period of viremia occurs in cats undergoing primary herpetic disease but not in cats undergoing recrudescent herpetic disease. These findings suggest that viremia may be important in the pathogenesis of primary herpetic disease but is less likely to contribute to the pathogenesis of recrudescent herpetic disease.
ABBREVIATIONS
| CPT | Cell preparation tube |
| CRFK | Crandell-Rees feline kidney |
| FHV-1 | Feline herpesvirus type 1 |
| pfu | Plaque-forming units |
| PID | Postinoculation day |
| SPF | Specific pathogen free |
| VI | Virus isolation |
IDEXX Laboratories, Sacramento, Calif.
Colorado State University Specialized Infectious Disease Laboratory, Fort Collins, Colo.
Lucy Whittier Molecular & Diagnostic Core Facility, Davis, Calif.
Kendall Monoject blood collection tubes, Tyco Healthcare Group, Mansfield, Mass.
DNeasy Tissue Kit, QIAGEN Inc, Valencia, Calif.
Polyester fiber–tipped/plastic applicator sterile swab, Fisher Scientific, Pittsburgh, Penn.
BD Vacutainer CPT, Becton, Dickinson & Co, Franklin Lakes, NJ.
Bright-Line hemacytometer, Hausser Scientific, Horsham, Penn.
Hydrocortisone-water soluble, Sigma-Aldrich Inc, St Louis, Mo.
DNA Micro Kit, QIAGEN Inc, Valencia, Calif.
References
- 1.
Bell SA, Balasuriya UB, Gardner IA, et al. Temporal detection of equine herpesvirus infections of a cohort of mares and their foals. Vet Microbiol 2006;116:249–257.
- 2.↑
Bell SA, Balasuriya UBR, Nordhausen RW, et al. Isolation of equine herpesvirus-5 from blood mononuclear cells of a gelding. J Vet Diagn Invest 2006;18:472–475.
- 3.
Bryans JT. On immunity to disease caused by equine herpesvirus 1. J Am Vet Med Assoc 1969;155:294–300.
- 4.
Goodman LB, Wagner B, Flaminio MJ, et al. Comparison of the efficacy of inactivated combination and modified-live virus vaccines against challenge infection with neuropathogenic equine herpesvirus type 1 (EHV-1). Vaccine 2006;24:3636–3645.
- 5.
Ch'ien LT, Whitley RJ, Nahmias AJ, et al. Antiviral chemotherapy and neonatal herpes simplex virus infection: a pilot study—experience with adenine arabinoside (ARA-A). Pediatrics 1975;55:678–685.
- 6.
Diamond C, Mohan K, Hobson A, et al. Viremia in neonatal herpes simplex virus infections. Pediatr Infect Dis J 1999;18:487–489.
- 7.↑
Schleiss MR, Bourne N, Bravo FJ, et al. Quantitative-competitive PCR monitoring of viral load following experimental guinea pig cytomegalovirus infection. J Virol Methods 2003;108:103–110.
- 8.↑
Nawynck HJ, Pensaert MB. Cell-free and cell-associated viremia in pigs after oronasal infection with Aujeszky's disease virus. Vet Microbiol 1995;43:307–314.
- 9.↑
Taus NS, Traul DL, Oaks JL, et al. Experimental infection of sheep with ovine herpesvirus 2 via aerosolization of nasal secretions. J Gen Virol 2005;86:575–579.
- 10.↑
Kaashoek MJ, Straver PH, Van Rooij EM, et al. Virulence, immunogenicity and reactivation of seven bovine herpesvirus 1.1 strains: clinical and virological aspects. Vet Rec 1996;139:416–421.
- 11.↑
Gaskell RM, Povey RC. Feline viral rhinotracheitis: sites of virus replication and persistence in acutely and persistently infected cats. Res Vet Sci 1979;27:167–174.
- 12.
Hoover EA, Rohovsky MW, Griesemer RA. Experimental feline viral rhinotracheitis in the germfree cat. Am J Pathol 1970;58:269–282.
- 13.↑
Nasisse MP, Guy JS, Davidson MG, et al. Experimental ocular herpesvirus infection in the cat. Sites of virus replication, clinical features and effects of corticosteroid administration. Invest Ophthalmol Vis Sci 1989;30:1758–1768.
- 14.
Reubel GH, Ramos RA, Hickman MA, et al. Detection of active and latent feline herpesvirus 1 infections using the polymerase chain reaction. Arch Virol 1993;132:409–420.
- 15.↑
Weigler BJ, Babineau CA, Sherry B, et al. High sensitivity polymerase chain reaction assay for active and latent feline herpesvirus-1 infections in domestic cats. Vet Rec 1997;140:335–338.
- 16.↑
Tham KM, Studdert MJ. Clinical and immunological responses of cats to feline herpesvirus type 1 infection. Vet Rec 1987;120:321–326.
- 17.
Karpas A, Routledge JK. Feline herpes virus: isolations and experimental studies. Zentralbl Veterinarmed [B] 1968;15:599–606.
- 18.
Van Pelt CS, Crandell RA. Pancreatitis associated with a feline herpesvirus infection. Compan Anim Pract 1987;1:7–10.
- 19.
Nasisse MP, Davis BJ, Guy JS, et al. Isolation of feline herpesvirus 1 from the trigeminal ganglia of acutely and chronically infected cats. J Vet Intern Med 1992;6:102–103.
- 20.
Kimura H, Futamura M, Kito H, et al. Detection of viral DNA in neonatal herpes simplex virus infections: frequent and prolonged presence in serum and cerebrospinal fluid. J Infect Dis 1991;164:289–293.
- 21.
Simons FA, Vennema H, Rofina JE, et al. A mRNA PCR for the diagnosis of feline infectious peritonitis. J Virol Methods 2005;124:111–116.
- 22.
van der Meulen KM, Favoreel HW, Pensaert MB, et al. Immune escape of equine herpesvirus 1 and other herpesviruses of veterinary importance. Vet Immunol Immunopathol 2006;111:31–40.
- 23.↑
Burgesser KM, Hotaling S, Schiebel A, et al. Comparison of PCR, virus isolation, and indirect fluorescent antibody staining in the detection of naturally occurring feline herpesvirus infections. J Vet Diagn Invest 1999;11:122–126.
- 24.
Clarke HE, Kado-Fong H, Maggs DJ. Effects of temperature and time in transit on polymerase chain reaction detection of feline herpesvirus DNA. J Vet Diagn Invest 2006;18:388–391.
- 25.
Holland JL, Outerbridge CA, Affolter VK, et al. Detection of feline herpesvirus 1 DNA in skin biopsy specimens from cats with or without dermatitis. J Am Vet Med Assoc 2006;229:1442–1446.
- 26.↑
Maggs DJ, Nasisse MP, Kass PH. Efficacy of oral supplementation with L-lysine in cats latently infected with feline herpesvirus. Am J Vet Res 2003;64:37–42.
- 27.↑
Stiles J, McDermott M, Willis M, et al. Comparison of nested polymerase chain reaction, virus isolation, and fluorescent antibody testing for identifying feline herpesvirus in cats with conjunctivitis. Am J Vet Res 1997;58:804–807.
- 28.
Sykes JE, Browning GF, Anderson G, et al. Differential sensitivity of culture and the polymerase chain reaction for detection of feline herpesvirus 1 in vaccinated and unvaccinated cats. Arch Virol 1997;142:65–74.
- 29.
Townsend WM, Stiles J, Guptill-Yoran L, et al. Development of a reverse transcriptase-polymerase chain reaction assay to detect feline herpesvirus-1 latency-associated transcripts in the trigeminal ganglia and corneas of cats that did not have clinical signs of ocular disease. Am J Vet Res 2004;65:314–319.
- 30.↑
Cullen CL, Wadowska DW, Singh A, et al. Ultrastructural findings in feline corneal sequestra. Vet Ophthalmol 2005;8:295–303.
- 31.
Lim CC, Cullen CL. Schirmer tear test values and tear film break-up times in cats with conjunctivitis. Vet Ophthalmol 2005;8:305–310.
- 32.↑
Cullen CL, Lim C, Sykes J. Tear film breakup times in young healthy cats before and after anesthesia. Vet Ophthalmol 2005;8:159–165.
- 33.↑
Maggs DJ, Clarke HE. Relative sensitivity of polymerase chain reaction assays used for detection of feline herpesvirus type 1 DNA in clinical samples and commercial vaccines. Am J Vet Res 2005;66:1550–1555.
- 34.
Nishiyama Y, Rapp F. Regulation of persistent infection with herpes simplex virus in vitro by hydrocortisone. J Virol 1979;31:841–844.
- 35.
Hoover EA, Griesemer RA. Experimental feline herpesvirus infection in the pregnant cat. Am J Pathol 1971;65:173–188.
- 37.↑
Hoover EA, Griesemer RA. Bone lesions produced by feline herpesvirus. Lab Invest 1971;25:457–464.
- 38.↑
Maggs DJ, Lappin MR, Reif JS, et al. Evaluation of serologic and viral detection methods for diagnosing feline herpesvirus-1 infection in cats with acute respiratory tract or chronic ocular disease. J Am Vet Med Assoc 1999;214:502–507.
- 39.↑
Bannasch MJ, Foley JE. Epidemiologic evaluation of multiple respiratory pathogens in cats in animal shelters. J Feline Med Surg 2005;7:109–119.
- 40.↑
Pedersen NC, Sato R, Foley JE, et al. Common virus infections in cats, before and after being placed in shelters, with emphasis on feline enteric coronavirus. J Feline Med Surg 2004;6:83–88.
