Pathogenesis of Experimental Rhesus Cytomegalovirus Infection

Abstract

Human cytomegalovirus (HCMV) establishes and maintains a lifelong persistence following infection in an immunocompetent host. The determinants of a stable virus-host relationship are poorly defined. A nonhuman primate model for HCMV was used to investigate virological and host parameters of infection in a healthy host. Juvenile rhesus macaques (Macaca mulatta) were inoculated with rhesus cytomegalovirus (RhCMV), either orally or intravenously (i.v.), and longitudinally necropsied. None of the animals displayed clinical signs of disease, although hematologic abnormalities were observed intermittently in i.v. inoculated animals. RhCMV DNA was detected transiently in the plasma of all animals at 1 to 2 weeks postinfection (wpi) and in multiple tissues beginning at 2 to 4 wpi. Splenic tissue was the only organ positive for RhCMV DNA in all animals. The location of splenic cells expressing RhCMV immediate-early protein 1 (IE1) in i.v. inoculated animals changed following inoculation. At 4 to 5 wpi, most IE1-positive cells were perifollicular, and at 25 wpi, the majority were located within the red pulp. All animals developed anti-RhCMV immunoglobulin M (IgM) antibodies within 1 to 2 wpi and IgG antibodies within 2 to 4 wpi against a limited number of viral proteins. Host reactivity to RhCMV proteins increased in titer (total and neutralizing) and avidity with time. These results demonstrate that while antiviral immune responses were able to protect from disease, they were insufficient to eliminate reservoirs of persistent viral gene expression.

Human cytomegalovirus (HCMV) infection does not generally result in clinical outcomes in immunocompetent hosts. Host antiviral responses rapidly restrict viral replication, such that most HCMV infections in healthy individuals are asymptomatic. While cellular and humoral antiviral immune responses can limit viral infection (4), HCMV can establish a lifelong persistence in a healthy host. Persistence is characterized by the presence of latent viral genomes that periodically reactivate to produce infectious virus (32). HCMV infects multiple cell types throughout the host, and recurrent virus can be shed from multiple sites (4, 29, 34). Rates of virus shedding can be as high as 39% in some study populations of healthy individuals (12, 14–16, 38, 41). Recurrent viral excretion is asymptomatic in most healthy hosts, although reactivated virus in immunosuppressed individuals presents serious clinical concern (4).

The mechanisms of HCMV persistence are not resolved. Results from in vitro and in vivo studies suggest that the virus utilizes a variety of strategies to maintain itself in a host despite vigorous cellular and humoral antiviral immune responses. Persistence strategies appear to include (i) broad cell tropism of the virus (36), (ii) differential patterns of gene expression in different cell types (17–19, 23, 24, 39, 40, 45, 49), (iii) novel patterns of gene expression (23, 24) and distinct genomic structures during latency (9), and (iv) expression of viral gene products that can modulate host immune responses (1, 2, 13, 20, 21, 28, 33, 37, 48). A major limitation to a better understanding of the determinants of a persistent infection is restricted knowledge concerning the early events of viral infection in vivo.

A nonhuman primate model of HCMV was used to investigate early stages of viral infection. Primate CMVs are closely related in terms of genetics (3, 8, 11, 25, 43), epidemiology (47), and the patterns of infection in immunocompetent and immunodeficient hosts (6, 27, 44, 47). In the study described here, juvenile rhesus macaques were experimentally inoculated with rhesus CMV (RhCMV) either intravenously (i.v.) or orally (p.o.), and virological and host parameters of infection were longitudinally analyzed.

MATERIALS AND METHODS

Animals and inoculations.

Healthy, RhCMV-seronegative, juvenile rhesus macaques (n = 9, 4 to 6 months old) were used for these studies. Animals were screened by Western blot analysis for seronegativity to RhCMV prior to inoculations. Animals were inoculated i.v. with 0.4 ml of virus stock that contained either 10⁶ (n = 3) or 10⁴ (n = 3) 50% tissue culture infectious doses of RhCMV strain 68-1 per ml. Three animals were inoculated p.o. with 10⁶ 50% tissue culture infectious doses of virus by placing the inoculum (0.4 ml) at the back of the tongue in anesthetized animals. The 68-1 strain of RhCMV has been demonstrated to be pathogenic in fetal macaques (44). Inoculated animals were monitored daily. Longitudinal blood samples were analyzed by complete blood counts and fluorescence-activated cell sorter analysis of CD3⁺-, CD4⁺-, and CD8⁺-cell populations (described below). Scheduled necropsies were performed at 2 (n = 1), 4 to 5 (n = 3), 8 (n = 1), 13 to 14 (n = 2), and 25 (n = 2) weeks postinfection (wpi). The inoculation and sampling schedule is detailed in Table 2.

TABLE 2.

Hematologic and histologic observations for inoculated macaques

Animal	Titer	Route	Necropsy (wpi)	Hematology	Histopathologya
29831	106	p.o.	2		LN, moderate LFH; Sp, moderate LFH
30457	106	p.o.	4		LN, mild to moderate LFH; tonsil, marked LFH; Sp, moderate LFH; Duod, mild lymphocytic-plasmacytic duodenitis
30460	106	p.o.	8		LN, mild to marked LFH; tonsil, marked LFH; Sp, moderate LFH; Duod, mild lymphocytic-plasmacytic duodenitis
29840	106	i.v.	4	Transient leukocytosis (4 wpi): mild neutrophilia, monocytosis, eosinophilia, moderate lymphocytosis	LN, moderate to marked LFH; Sp, moderate to marked LFH, sinus neutrophilia and lymphocytosis; tonsil, tonsilitis, neutrophilic, marked LFH; Jej/Duod, dilated lacteal; Ki, mild membranoproliferative glomerulopathy
29721	104	i.v.	4		LN, mild LFH; tonsil, tonsilitis, neutrophilic, marked LFH; Sp, sinus neutrophilia; Th, mild lymphoid depletion; Duod/Jej, mild LFH; Duod/Il, mild LFH; Li, moderate centrolobular vacuolar degeneration; Ki, mild membranoproliferative glomerulopathy; Lu, prominent pulmonary lymphoid tissue, focal, perivascular lymphocytosis
29740	104	i.v.	12	Mild monocytosis (2, 9 wpi), mild eosinophilia (7, 9 wpi)	LN, marked LFH, hyalinization of germinal centers; Sp, marked LFH, hyalinization of germinal centers, sinus neutrophilia and lymphocytosis; tonsil, marked LFH; Duod/Jej, marked LFH; Lu, prominent lymphoid tissue, multifocal
29762	104	i.v.	12	Transient leukocytosis (5, 9 wpi): mild neutrophilia, monocytosis, moderate lymphocytosis	LN, moderate to marked LFH, sinus histiocytosis; Sp, moderate LFH, sinus neutrophilia; tonsil, tonsilitis, neutrophilic, marked LFH; Duod, marked mucosal LFH; Il, marked submucosal LFH; Li, mild multifocal lymphocytic hepatopathy
29848	106	i.v.	25	Transient leukocytosis (8 wpi): mild lymphocytosis, mild monocytosis (6, 8, 22 wpi), marked eosinophilia	LN, moderate LFH, sinus histiocytosis; Sp, marked LFH; Duod, mild LFH; BM, mild myeloid hyperplasia
29850	106	i.v.	25	Persistent leukocytosis (2, 3, 6, 8, 17 wpi): mild neutrophilia (3, 22, 25 wpi), mild lymphocytosis (3, 8, 13, 17 wpi), mild monocytosis (1, 2, 3, 6, 8, 17, 22 wpi), mild eosinophilia (2, 8 wpi)	LN, moderate to marked LFH; tonsil, moderate LFH; Sp, moderate LFH sinus neutrophilia; Duod/Jej, moderate LFH; Duod, enteritis, plasmacytic, diffuse, crypts; Li, moderate centrolobular vacuolar degeneration; Lu, prominent pulmonary lymphoid tissue; BM, moderate myeloid hyperplasia

^aLFH, lymphofollicular hyperplasia; Sp, spleen; Duod, duodenum; Jej, jejunum; Ki, kidney; Th, thymus; Il, ileum; Li, liver; Lu, lung; BM, bone marrow. 

Hematologic evaluation and CD4/CD8 T-lymphocyte immunophenotyping.

Complete blood counts were performed by the Clinical Laboratory at the California Regional Primate Research Center with EDTA-anticoagulated blood. Fifty microliters of whole blood was incubated for 15 to 30 min at room temperature with monoclonal antibodies specific for CD3 (10 μl, fluorescein isothiocyanate conjugated) (catalogue no. 35694X; Pharmingen, San Diego, Calif.), CD4 (5 μl, phycoerythrin conjugated) (catalogue no. 703430; Ortho Diagnostics, Raritan, N.J.), and CD8 (5 μl, peridinin chlorophyll protein [PerCP] conjugated) (catalogue no. 347314, Becton Dickinson, Mountain View, Calif.). Samples were processed (Q-Prep; Coulter, Hialeah, Fla.) and analyzed by three-color flow cytometry with a FACScan (Becton Dickinson). Fluorescence-activated cell sorter analysis was performed by the Optical Biology Laboratory, Department of Medical Pathology, University of California, Davis.

Necropsy and histology.

Animals were culled at defined time points during the study period (see Table 2). Complete necropsy examinations were performed. Tissues were utilized for histologic examination and nucleic acid purification. Tissues were fixed in 10% formalin for a period of time not exceeding 5 days prior to paraffin embedding (46).

PCR.

DNA was purified from necropsy tissues by using a lysis buffer containing 1% sodium dodecyl sulfate, 20 mM Tris (pH 7.5), 250 mM NaCl, 20 mM EDTA, and 0.5 mg of proteinase K per ml. Tissues were incubated in lysis buffer overnight at 63°C, and fresh lysis buffer was added until the tissues were completely digested. After phenol-chloroform extraction and precipitation with ethanol, DNA concentrations were determined spectrophotometrically. Viral DNA was purified from 200 μl of plasma by using the QIAmp blood kit (Qiagen, Valencia, Calif.); DNA was eluted in a volume of 50 μl of water. Nested PCR (40 cycles of 1 min at 94°C, 1 min at 55°C, and 1 min at 72°C) was performed with 1 μg of DNA as the template and primers to exon 5 of immediate-early protein 2 (IE2) (8). For the first round of PCR, primers PAB196 (5′ GCCAATGCATCCTCTGGATGTATTGTGA 3′) and PAB249 (5′ TGCTTGGGGAATCTCTGCAC 3′) were used. For the second nested round of PCR, 5 μl of the first-round product of PCR was amplified by using primers PAB201 (5′ CCCTTCCTGACTACTAATGTAC 3′) and PAB248 (5′ TTGGGGAATCTCTGCACAAG 3′). A volume of 10 μl of plasma DNA eluate was used as the template for PCR.

Immunohistochemistry.

The expression and distribution of RhCMV IE1 were determined by immunohistochemistry. Briefly, slides were deparaffinized for 20 min at 65°C and rehydrated (46). They were then incubated in 3% hydrogen peroxide–distilled water for 10 min, followed by incubation in 10 mM sodium citrate for 4 h at 90°C. Slides were cooled to 25°C for 20 to 30 min and then washed three times in phosphate-buffered saline (PBS). To reduce nonspecific binding, tissue sections were treated for 30 min with Protein Block Serum-Free (Dako, Carpinteria, Calif.) at 25°C. After being washed with PBS, slides were incubated with primary antibody (1:3,200 in PBS–0.1% Tween 20 [PBS-T]) overnight at 4°C. For these studies, a rabbit polyclonal antiserum was generated against a bacterially synthesized protein corresponding to exon 4 of the RhCMV IE1 gene (8) (proteins were synthesized by Casey Morrow, University of Alabama at Birmingham). Following three washes in PBS-T, secondary antibody (biotinylated goat anti-rabbit, 1:800; Vector, Carpinteria, Calif.) was added and left for 1 h at 25°C. The slides were washed again three times in PBS-T, and avidin-biotin complex–peroxidase (ABC) (Vector) was added, followed by diaminobenzidine (DAB) (Vector) as a substrate for 1 h at 25°C. Tissues were further processed according to published protocols (46).

For double-immunolabeling experiments, tissues were incubated overnight at 4°C with antisera against both IE1 and markers specific for B cells (CD20; 1:500 dilution) (Dako; catalogue no. M0755), macrophages (HAMS56; 1:400) (Dako; M0634), dendritic and endothelial cells (p55/fascin; 1:5,000) (Dako; M3567), or plasma cells (immunoglobulin G [IgG]; 1:200) (Accurate Chemical Co., Westbury, N.Y.; YNGM01GGFCP). The antibody against fascin recognizes both endothelial and dendritic cells within the spleen and other lymphoid organs (35). After being washed three times in PBS-T, tissues were incubated with biotinylated secondary antibody (1:800) for 1 h at 25°C and then incubated with ABC, followed by DAB staining (as described above). After optimal color development, slides were incubated in distilled water to stop additional color development and then in 3% hydrogen peroxide (10 min at 25°C) to inactivate the conjugated peroxidase. After being washed in PBS-T, tissues were incubated in biotinylated anti-rabbit antibody (1:800), ABC, and VIP peroxidase substrate (Vector) to yield a purple product in IE1-expressing cells. Because the anti-T (CD3; 1:400) (Dako; A0452) antibody was rabbit polyclonal antiserum, double-labeling experiments for IE1 and CD3 were not feasible. Four-micrometer serial tissue sections were individually analyzed for IE1 or CD3 expression and compared.

Western blot analysis.

Western blot analysis for anti-RhCMV IgG was performed by previously published protocols (47) with a 1:100 dilution of plasma and RhCMV-infected rhesus skin fibroblasts as the antigen.

Neutralization assays.

Assays to measure neutralizing antibody titers were performed as described by Andreoni et al. (5) with modifications. Primary rhesus skin fibroblasts were plated at a density of 2 × 10⁴ cells per well of a 96-well plate on the day prior to the neutralization assay. Heat-inactivated (30 min, 56°C) plasma was twofold serially diluted in Dulbecco modified Eagle medium–10% calf serum in a final volume of 24 μl. A stock of RhCMV of known titer was diluted to give a final titer of 100 infectious virus particles per 50 μl; 168 μl of diluted virus was added to each plasma dilution. Samples were vortexed and incubated at 37°C in a water bath for 2 h, and then 50 μl of plasma-virus was added in triplicate to cells and incubated at 37°C in a CO₂ incubator for 3.5 h. Virus was removed, and cells were fed normal growth medium. Sixteen hours later, the cells were fixed in methanol-acetone (1:1) for 20 min, washed four times in PBS, and then incubated in 1% bovine serum albumin (BSA) in PBS-T (BSA–PBS-T) for 2 h at 25°C. The blocking solution was removed, and anti-IE1 rabbit polyclonal antibody (diluted 1:3,200 in BSA–PBS-T) was added and left for 2 h at 25°C. Plates were washed three times in PBS-T and then incubated with biotinylated goat anti-monkey IgG (1:800) (Sigma, St. Louis, Mo.) for 1 h. After being washed, the plates were incubated in ABC Elite (Vector) for 30 min, followed by addition of DAB chromogenic substrate. The number of stained nuclei in each well was counted and compared with those in wells incubated with virus and medium alone. The neutralization titer was calculated as the inverse of the plasma dilution resulting in a 50% reduction of IE1-positive cells per well.

ELISA antiviral antibody titers.

Anti-RhCMV IgM and IgG antibodies were quantified by enzyme-linked immunosorbent assay (ELISA), using a modification of published protocols (31). Briefly, primary rhesus skin fibroblasts were infected with RhCMV and harvested in NaOH-glycine buffer when the cells exhibited 100% cytopathic effect. After sonication of the lysate, the protein concentration was determined, and aliquoted extracts (500 μg/ml) were stored at −20°C. Individual wells of a 96-well plate were coated with 1 μg of protein in carbonate buffer (pH 9.6) (31) overnight at 4°C. After being washed three times in PBS-T, wells were incubated with blocking solution (BSA-PBS) for 2 h at 25°C. Plasma samples, including a pool of RhCMV-seronegative plasma, were serially diluted in BSA–PBS-T. The wells were washed, and diluted plasma samples were loaded in duplicate at 100 μl/well. Wells were incubated with plasma for 2 h at 25°C, washed three times, and incubated with 100 μl of goat anti-monkey IgG peroxidase-conjugated antibody (diluted in BSA–PBS-T) (Kirkegaard and Perry Laboratories, Gaithersburg, Md.) for 1 h at 25°C. Peroxidase-conjugated goat anti-monkey IgM antibody (Kirkegaard and Perry) (diluted in BSA–PBS-T) was used for the IgM ELISA. The tetramethylbenzidine liquid substrate system (Sigma) was used as a substrate (30 min at 25°C), and the reaction was stopped with 0.5 M H₂SO₄ (50 μl/well). Absorbency at 450 nm (A₄₅₀) was recorded within 1 h.

Antibody avidity.

Antibody avidity was determined by published protocols (10) with modifications. Briefly, the first plasma sample with detectable anti-RhCMV IgG and the terminal plasma sample were diluted 1:100 in BSA–PBS-T. Following incubation and removal of diluted plasma, the wells were incubated for 15 min with increasing concentrations (0, 1.0, 1.5, 2.0, 2.5, and 3.0 M) of sodium thiocyanate (NaSCN) diluted in PBS. The wells were subsequently washed and processed as described above. The avidity index was calculated as the molar concentration of NaSCN required to reduce the A₄₅₀ by 50% compared to that observed in the absence of NaSCN.

RESULTS

Hematologic analysis and T-lymphocyte immunophenotyping.

All animals inoculated with RhCMV remained clinically healthy during the study. None of the p.o. inoculated animals demonstrated any significant hematologic change from preinoculation values. However, all six monkeys inoculated i.v. exhibited hematologic abnormalities in association with viral infection. Four animals (29850, 29840, 29762, and 29848) developed leukocytosis at 2, 4, 5, and 8 wpi, respectively, compared to reference ranges; leukocytosis was transient, except for animal 29850, which exhibited persistent leukocytosis 2 to 8 wpi. Monocytosis was periodically observed in five of six animals. Monocytosis was particularly noteworthy in animal 29840 at 4 wpi; it coincided with a marked neutrophilia (14,452/μl) and lymphocytosis (22,684 lymphocytes/μl). Transient neutrophilia was also observed in animals 29762 and 29850 (at 5 and 3 wpi, respectively). Erythrocyte and platelet numbers remained within the reference ranges for the study period for all animals except 29840. At 4 wpi, this animal exhibited a marked thrombocytopenia (16 × 10⁵/μl).

Viral infection was associated with acute changes in CD4/CD8 ratios in all i.v. inoculated animals. No changes were observed in p.o. inoculated animals. For i.v. inoculations, an initial decrease in CD4/CD8 ratios was seen at 2 wpi, followed by a gradual return to preinoculation values beginning at 3 wpi. The most prominent changes in ratios were observed for animals 29840 and 29848, with CD4/CD8 ratios dropping from 2.1 and 2.3, respectively, to 0.7. Decreases in CD4/CD8 ratios were due to increases in both CD8⁺ CD3⁺ T lymphocytes and CD8⁺ CD3⁻ natural killer (NK) cells. No consistent changes were observed for absolute CD4⁺-cell numbers.

Immunologic analysis.

All animals except 29831 (inoculated p.o.) developed detectable antiviral IgM antibodies by 2 wpi (Table 1). Peak titers were observed at 1 to 2 wpi and ranged from 640 to >5,120. Titers of IgM antibodies decreased rapidly and were generally undetectable by 4 to 8 wpi; low levels of IgM antibodies were noted in animal 29850 at 13 and 25 wpi.

TABLE 1.

Antiviral immunological parameters in RhCMV-inoculated juvenile macaques

Inoculation	Animal	Titera (wpi)					Avidity indexd (wpi)
		Peak IgM	IgG		NAb
			Firstb	Lastc	First	Last	First	Last
p.o.	29831	NDe	ND (2)
	30457	640 (2)	200 (2)	100 (4)		16 (4)		0.7 (4)
	30460	640 (2–4)	1,600 (4)	1,600 (8)	128 (4)	512 (8)	0.8 (4)	1.4 (8)
i.v.	29840	640 (2)	800 (2)	400 (4)	128 (4)		0.8 (2)	0.9 (4)
	29721	1,280 (2)	400 (4)					1.1 (5)
	28740	2,560 (2)	200 (2)	800 (13)	16 (2)	512 (13)	1.1 (2)	2.0 (13)
	29762	640 (2)	400 (5)	800 (13)			0.8 (5)	1.8 (13)
	29848	1,280 (1)	200 (2)	6,400 (25)	32 (2)	2,048 (25)	0.7 (2)	3.0 (25)
	29850	>5,120 (1–2)	200 (2)	6,400 (25)			0.8 (2)	2.8 (25)

^aTiter is defined as the reciprocal of the dilution producing an A₄₅₀ value greater than the mean sum for a pool of seronegative plasma plus 0.1 absorbency unit. Minimum dilutions: IgM, 1:40; IgG, 1:100; neutralization antibody (NAb), 1:16. 

^bFirst time point with positive IgG titer. 

^cLast time point (i.e., necropsy) with positive IgG titer. 

^dMolar concentration of NaSCN required to reduce the initial A₄₅₀ by 50% (1:100 dilution). 

^eND, not detected. 

Anti-RhCMV IgG antibodies were first detected at 2 wpi in five animals (one inoculated p.o. and four inoculated i.v.) (Table 1) and at 4 wpi in three animals; there were no detectable IgG titers for animal 29831 (p.o.) at the 2 wpi necropsy time point (100-fold minimal antibody dilution). Antiviral IgG antibodies reached maximal titers between 10 and 25 wpi for the two animals (29848 and 29850) studied for 25 weeks (Table 1). Recent studies have suggested that antibody avidity is a qualitative measure of protection (10). Antibody avidity was determined for each animal (except 29831) by using the first IgG-positive plasma sample and the terminal plasma sample. For each animal with two time points analyzed, antibody avidity increased with time. Increases were observed at early time intervals (animal 29840 at 2 to 4 wpi), and avidity continued to increase until at least 25 wpi (animals 29848 and 29850) (Table 1). It should be noted that anti-RhCMV titers for animals 29848 and 29850 at 25 wpi were comparable to those observed in seropositive adult macaques housed in outdoor breeding corrals (data not shown). However, the avidity indices for animals 29848 and 29850 were at the lower end of the range observed in corral animals (avidity index of 3.0 to 5.0) (data not shown), indicating that the time frame for antibody maturation is greater than 6 months.

The kinetics of antiviral antibody development coincided with the appearance of neutralizing antiviral antibodies. Microneutralization assays (5) were performed with longitudinal plasma samples from p.o. inoculated animals 30457 and 30460 and i.v. inoculated animals 29840, 29740, and 29848. Neutralizing antibody titers were first detected at 2 wpi for animals 29740 and 29848 and at 4 wpi for the other three animals examined (Table 1). Titers of neutralizing antibodies increased until 13 weeks for animals 29740 and 29848.

Host immune responses were directed initially against a limited number (one to seven) of viral proteins with approximate sizes of 49, 60, 65, 80, 86/87, and 105 kDa; representative examples are presented for animals 30460 (inoculated p.o.) and 29850 (inoculated i.v.) in Fig. 1. The pattern of immune reactivity became increasingly complex at later stages of infection, particularly for animals 29848 (not shown) and 29850. The relative intensities against some proteins (e.g., the 85/87-kDa protein) appeared to increase during the course of observation, while others (e.g., the 49-kDa protein) stabilized quickly. Other proteins, particularly the 60-, 65-, and 80-kDa proteins, appeared to decline in relative titer. The identities of reactive proteins are not known, although antibodies to glycoprotein B were detected at 2 to 3 wpi (10a).

FIG. 1.

Western blot analysis of plasma from inoculated animals. Longitudinal plasma samples from p.o. (30460) and i.v. (29840, 29740, and 29848) inoculated animals were evaluated by Western blotting for reactivity to RhCMV antigens. The time points for each plasma sample (wpi) are given below each lane. Molecular masses are shown at the right and left.

Detection of RhCMV DNA in plasma.

RhCMV genomic DNA was amplified from DNA purified from plasma by using primers to exon 5 of IE2 (8). RhCMV DNA was first detectable in plasma at 1 wpi for all animals except 30457 (Fig. 2). For animal 30457, RhCMV DNA was detected initially in plasma at the 2 wpi time point. Four animals, one inoculated p.o. (30457) and three inoculated i.v. (29762, 29848, and 29850), were PCR positive for RhCMV DNA in plasma at more than one time point, although the amplified products were barely visible by ethidium bromide staining after the 1 wpi time point (Fig. 2). No attempt was made to culture virus from plasma. Therefore, it is not known if RhCMV DNA in plasma represented infectious virus.

FIG. 2.

PCR analysis of RhCMV DNA in inoculated animals. Longitudinal plasma samples (top panels) and tissues obtained at necropsy (bottom panels) from p.o. (left panels) and i.v. (right panels; 29840, 29740, and 29848 only) inoculated (Inoc.) animals were analyzed for RhCMV DNA. For plasma samples, wpi are shown (week 0 was not available for animals 29831 and 29762). Samples with a positive signal (+) are indicated. Tissue samples analyzed were axillary LN (lanes 1), bone marrow (lanes 2), pancreas (lanes 3), ileum (lanes 4), inguinal LN (lanes 5), jejunum (lanes 6), kidney (lanes 7), lung 1 (lanes 8), lung 2 (lanes 9), submandibular gland (lanes 10), spleen (lanes 11), thymus (lanes 12), tonsil (lanes 13), liver (lanes 14), parotid gland (lanes 15), and esophagus (p.o. inoculated animals only) and duodenum (i.v. inoculated animals only) (lanes 16). Lane P, positive control tissue; lane M, molecular weight markers.

Histopathology.

The most significant histopathologic finding was lymphofollicular hyperplasia in all animals, involving the duodenum, jejunum, lymph nodes (LN), spleen, and tonsils (Table 2). Lymphoid hyperplasia ranged from mild to marked or severe and was noted at both early and late time points; hyperplasia was most prominent in animals 29740 (inoculated i.v.) and 30460 (inoculated p.o.) and was accompanied by germinal center hyalinization. In addition, a neutrophilic splenitis occurred in two p.o. inoculated animals (30457 and 30460) and all i.v. inoculated animals and was characterized by prominent neutrophilic infiltrates in the red pulp. Splenitis ranged from mild (animals 29848 and 29721) to moderate (30457, 29840, and 29762) to marked (30460, 29740, and 29850). Viral inclusions were not observed in any tissue.

Immunohistochemistry.

Immunohistochemical analysis for IE1 expression was performed with tissues that were positive for RhCMV DNA (discussed below). IE1 expression was detected only in the spleens of i.v. inoculated animals and was localized to the nucleus of splenic cells (Fig. 3). The cellular location of IE1-positive cells within the spleen changed during the course of infection. At 4 to 5 wpi, most cells expressing IE1 were perifollicular (Fig. 3A). Lower numbers of IE1-positive cells were seen within the follicles and in the red pulp. To estimate the extent of viral expression within the spleen, the number of IE1-positive follicles was counted. A total of 86% (68 of 79) of the splenic follicles in animal 29840 and 14% (14 of 100) in animal 29721 contained cells staining for IE1 in a representative section (data not shown). In contrast, the majority of splenic cells expressing IE1 at 25 wpi (Fig. 3C) were located within the red pulp. Only occasional IE1-positive cells were present in perifollicular areas; no follicular germinal centers were positive for IE1. For animal 29850, 14% of the perifollicular areas (14 of 99) had cells expressing IE1; 2% of the perifollicular areas (2 of 93) in animal 29848 were positive for IE1 expression. The two animals analyzed at 12 to 13 wpi (29740 and 29762) exhibited an intermediate pattern, with IE1-positive cells in both perifollicular areas and the red pulp (Fig. 3B).

FIG. 3.

Immunohistochemical localization of RhCMV IE1-expressing cells in the spleen. (A to C) Spleen sections from animals 29840 (A), 29740 (B), and 29848 (C) were assayed for RhCMV IE1 expression. IE1-positive cells are brown (DAB), and the cells are counterstained with hematoxylin (blue). The location of a representative follicle (f) is indicated. (D and E) Cells were double labeled for IE1 expression and markers for endothelial cells (D) or macrophages (E). (D) Endothelial cells are brown, and IE1-positive cells are purple. (E) Macrophages are pink, and IE1-positive cells are brown. Double-labeled cells are indicated by arrows. Many of the IE1-expressing cells are not stained with either cell type marker.

Histologically, it appeared that multiple cell types supported viral gene expression (Fig. 3A to C). Cells positive for RhCMV IE1 expression within the mantle zone of the follicle usually contained a thin, flattened nucleus and cytoplasm, suggestive of endothelial or fibroblast cells. Positive cells either within the follicle or in the parenchyma of the red pulp had variable morphologies. Many of these cells contained either a large round or a reniform nucleus (consistent with that of a macrophage), while others were elliptical or irregular in shape.

To further examine the cell tropism for RhCMV in the spleen, double immunohistochemistry was performed with antibodies to RhCMV IE1 and cell-specific markers for B, T, macrophage, endothelial and dendritic, or plasma cells. The results demonstrated that IE1 was occasionally colocalized to endothelial cells within both perifollicular areas (data not shown) and the sinusoids of the red pulp (Fig. 3D) and to tissue macrophages in the red pulp (Fig. 3E). However, most IE1-positive cells did not costain for either the macrophage or endothelial cell marker; representative IE1-positive cells not stained with either the macrophage or endothelial cell marker are seen in Fig. 3D and E. In addition, there was no evidence for expression of IE1 within the other cell types examined (plasma, B, or dendritic) in any animal (data not shown). Analysis of serial splenic sections for IE1 and the CD3 T-cell marker also failed to conclusively identify IE1 expression within this cell type.

Detection of RhCMV DNA in tissues.

The spleen was the only tissue that was PCR positive in all nine animals (Fig. 2). For i.v. inoculations, axillary LN were positive in five animals, inguinal LN were positive in four, bone marrow was positive in three, and liver was positive in two. Multiple tissues (pancreas, ileum, kidney, lung, tonsil, thymus, submandibular gland, and mesenteric LN) were PCR positive in only one animal. These same tissues were positive for RhCMV DNA in at least one of the p.o. inoculated animals, except for inguinal and mesenteric LN. Other than the consistent positive signal observed for the spleen, there was no apparent pattern to the particular tissues or number of tissues positive for RhCMV DNA at the different time points analyzed. The limit of sensitivity of PCR detection of viral DNA was approximately 10 viral genome equivalents per μg of cellular DNA, or 10 viral genome equivalents per 3 × 10⁵ cells (data not shown).

DISCUSSION

Experimental inoculation of rhesus macaques with RhCMV paralleled two fundamental aspects of HCMV infection in humans: (i) limited pathogenic potential in healthy, immunocompetent hosts and (ii) persistent gene expression in the presence of host antiviral immune responses. The mechanisms for these two characteristic features of CMV infection were not identified. Our observations indicated that the course of viral infection was not a function of the route of RhCMV inoculation and that establishment of a stable virus-host relationship probably required many months.

Virological and host parameters of infection were similar after either parenteral or mucosal inoculation. Infection was characterized by an initial burst of RhCMV DNA in the plasma at 1 to 2 wpi in all animals. RhCMV disseminated to a common, but not identical, spectrum of tissues by the earliest time point examined for each inoculation method (2 and 4 wpi for p.o. and i.v. inoculation, respectively). The tissues positive for RhCMV DNA were consistent with those identified as sites for HCMV (36), including those, such as bone marrow, identified as possible reservoirs of latent virus (19). The early targeting of the macrophage and endothelial cells within the spleen by RhCMV has been observed for murine CMV (MCMV) (30, 42). PCR detection of RhCMV DNA in plasma and tissues suggests that after inoculation there is an initial amplification of RhCMV at the primary site of infection followed by hematogenous spread of the virus at 1 to 2 wpi. The virus then spread to multiple tissues by 2 to 4 wpi, the earliest time points examined for p.o. and i.v. inoculation, respectively. The observation that DNA was observed in plasma in both p.o. and i.v. inoculated animals indicates that RhCMV dissemination from the primary site of infection to the tissues required a blood-borne intermediate step. An alternative model might have been that direct inoculation of virus into the circulation obviated a hematogenous phase.

Host responses to RhCMV followed similar kinetics after either p.o. or i.v. inoculation. All animals except 29831 developed antiviral antibodies within 2 weeks of inoculation, with increases in titers (total and neutralizing) and avidity indices for at least 8 and 25 wpi (for p.o. and i.v. inoculation, respectively). Antiviral immune responses in immunocompetent animals were sufficient to prevent RhCMV disease. Inoculation of fetal macaques or juvenile macaques coinfected with simian immunodeficiency virus with similar titers of RhCMV can result in a rapid, severe pathogenic outcome (reference 44 and unpublished data). The absence of both disease and prolonged presence of DNA in plasma suggested that early immune responses effectively restricted RhCMV sequelae. The decline in plasma DNA levels occurred when anti-RhCMV IgG antibodies were low (or undetectable) in titer and of low avidity. This argues that IgM and innate immune responses were important for limitation of acute disease but were not effective for preventing viral dissemination throughout the host. The kinetics of cell-mediated antiviral immunity were not evaluated in this study. Host immune responses were insufficient to eliminate reservoirs of persistent virus gene expression. Prolonged (i.e., 6 months) RhCMV gene expression (IE1) was observed in the spleens of i.v. inoculated animals that had developed high-avidity and high-titer neutralizing antiviral antibodies. It was not determined whether gene expression was associated with the production of infectious virus.

An intriguing aspect of these studies was the differential localization in the spleen of IE1-positive cells at different times following i.v. inoculation. Assuming that the two i.v. inoculated animals examined at each of the three time points represented a continuum of the infection process, there was a pronounced change in the sites of IE1-expressing cells. This shift was noted for a change from a predominantly follicular expression pattern at 4 wpi to one of predominantly red pulp expression at 25 wpi. The transition for the location of IE1-positive cells has not been previously described. The localization of virally infected cells within and around the follicles at 4 wpi in this study was similar to that observed with mice experimentally inoculated with MCMV (42). In that study, the authors postulated that the pattern of MCMV-infected cells corresponded to that of marginal-zone macrophages and dendritic cells. The morphological features of IE1-expressing cells in the macaques were consistent with multiple cell types, including macrophages, endothelial cells, and other cells. We colocalized IE1 expression with markers for endothelial cells in the follicles (Fig. 3D) and tissue macrophages (Fig. 3E) in the red pulp. However, the vast majority of IE1-expressing cells were not colabeled with these cell markers or with those for dendritic, B, plasma, or T cells. The identity of these IE1-expressing cells remains unknown, and their characterization is important, since they constituted the bulk of virus-positive cells within the spleen.

Several non-mutually exclusive theories can describe the apparent shift in virus expression within the spleen. One is that the pattern of IE1-positive cells represents trafficking of virus through the spleen at different stages of infection. According to this hypothesis, infection of follicle-associated cells during the acute phase is necessary for subsequent infection of cells within the red pulp. Alternatively, there may be differential susceptibility of cells within the spleen to host immune mechanisms. The host might preferentially eliminate virus-expressing cells within the follicle, such that IE1-positive cells within the red pulp predominated by 25 wpi. Finally, viral gene expression may decrease to undetectable levels in follicle-associated cells prior to doing so in cells in the red pulp. Such a mechanism can be considered an extension of the second hypothesis, particularly if virus-expressing cells in the follicle are immunologically more susceptible than those in the red pulp. The temporal profile of RhCMV IE1-positive cells within the spleen implies a continuous process in the establishment of a persistent infection.

In summary, conditions have been established for inoculation of healthy rhesus macaques with RhCMV to study viral and host parameters of infection. The results of this study and others (3, 7, 8, 22, 25, 26, 43, 44, 47) demonstrate that the nonhuman primate system is an extremely relevant model for the study of HCMV persistence and pathogenesis.