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Journal of Virology, July 2008, p. 6524-6535, Vol. 82, No. 13
0022-538X/08/$08.00+0     doi:10.1128/JVI.00502-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Human Cytomegalovirus Secretome Contains Factors That Induce Angiogenesis and Wound Healing

Jerome Dumortier,⁴ Daniel N. Streblow,⁴ Ashlee V. Moses,⁴ Jon M. Jacobs,⁵ Craig N. Kreklywich,² David Camp,⁵ Richard D. Smith,⁵ Susan L. Orloff,^1,2,3 and Jay A. Nelson^2,4*

Veterans Affairs, Portland VAMC, Portland, Oregon,¹ MMI,² Department of Surgery, Oregon Health Sciences University, Portland, Oregon,³ Vaccine and Gene Therapy Institute, Oregon Health Sciences University, Portland, Oregon,⁴ Biological Sciences Division, Pacific Northwest National Laboratory, Richland, Washington 99352⁵

Received 6 March 2008/ Accepted 21 April 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Human cytomegalovirus (HCMV) is implicated in the acceleration of a number of vascular diseases including transplant vascular sclerosis (TVS), the lesion associated with chronic rejection (CR) of solid organ transplants. Although the virus persists in the allograft throughout the course of disease, few cells are directly infected by CMV. This observation is in contrast to the global effects that CMV has on the acceleration of TVS/CR, suggesting that CMV infection indirectly promotes the vascular disease process. Recent transcriptome analysis of CMV-infected heart allografts indicates that the virus induces cytokines and growth factors associated with angiogenesis (AG) and wound healing (WH), suggesting that CMV may accelerate TVS/CR through the induction and secretion of AG/WH factors from infected cells. We analyzed virus-free supernatants from HCMV-infected cells (HCMV secretomes) for growth factors, by mass spectrometry and immunoassays, and found that the HCMV secretome contains over 1,000 cellular proteins, many of which are involved in AG/WH. Importantly, functional assays demonstrated that CMV but not herpes simplex virus secretomes not only induce AG/WH but also promote neovessel stabilization and endothelial cell survival for 2 weeks. These findings suggest that CMV acceleration of TVS occurs through virus-induced growth factors and cytokines in the CMV secretome.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Numerous epidemiological and animal studies link human cytomegalovirus (HCMV) to the acceleration of vascular diseases including arterial restenosis, atherosclerosis, and transplant vascular sclerosis (TVS) (23, 24, 37). Recent advances in transplantation have significantly impacted short-term allograft and patient survival; however, long-term graft survival has not improved, due largely to chronic rejection (CR). The prevalence of CR is a concern, since retransplantation is the sole effective therapy. TVS represents the hallmark of CR in vascularized solid organ transplants, and HCMV infection nearly doubles the 5-year rate of cardiac graft failure due to accelerated TVS (11). In heart transplant recipients, ganciclovir, a potent inhibitor of CMV replication, delays the time to allograft rejection (25, 44). A higher incidence of viral DNA in the explant vascular intima from patients with cardiac allograft TVS than in explants without vasculopathy further underscores the influence of HCMV on CR development (47). In kidney transplant patients, the presence of HCMV infection, whether asymptomatic or displaying overt symptoms, negatively impacts allograft survival (9). The role of HCMV in TVS/CR development is clear; however, the mechanisms involved in this process remain illusive for the following reasons: HCMV disease etiology is multifactorial; HCMV is ubiquitous throughout the human population; HCMV infection is lifelong and infects all of the cell types involved in TVS, including smooth muscle cells (SMC), endothelial cells (EC), and macrophages; and HCMV evades the immune system by remaining latent, and clinically silent reactivation is difficult to detect (3, 18, 19, 28, 30). Animal models provide an ideal tool for studying the association between CMV and TVS and have confirmed that herpesvirus infections play a role in vascular disease. Acute infection with rat (R)CMV accelerates TVS in rat organ transplantation, leading to graft failure (29, 35, 38). Similar to the clinical scenario, antiviral therapy reduces the acceleration of rejection, underscoring the need for viral replication (42, 48). Importantly, the effects of RCMV on TVS acceleration are not organ type specific but occur in various solid organs (29, 35, 38, 43).

TVS is characterized histologically by diffuse concentric intimal proliferation resulting in vessel stenosis and ultimately allograft failure (1). A few macrophages, T cells, NK cells, and B cells are seen in early lesions, while late lesions are associated with a thickened intima containing SMC interspersed with macrophages (6). Activated inflammatory cells and SMC within and near vascular lesions are important local sources of proangiogenic factors (2). TVS development involves chronic perivascular inflammation, EC dysfunction, and SMC migration/proliferation, resulting in neointimal thickening of the allograft arterial wall (1, 12, 22). A growing body of evidence supports a role for angiogenesis (AG) and wound healing (WH) in the development of vascular diseases (17, 34). Importantly, pathological processes involved in TVS and other vascular diseases parallel many cellular events that mediate normal AG/WH. Complex interactions between cells and surrounding regulatory factors lead to cellular migration, proliferation, and tissue remodeling.

AG is broadly divided into three phases: vessel destabilization, proliferation/migration, and vessel maturation (4). In healthy humans, AG is restricted to the formation of placental and endometrial tissue, hair follicle vascularization, and WH due to positive and negative regulatory factors. Where vessel growth is required, the regulatory balance tips toward proangiogenic factors. Increasing AG inhibitors and vessel stabilization factors achieves restoration of a steady state. Breakdown of the tightly regulated angiogenic balance contributes to a variety of pathological disorders, including cancer and autoimmune and cardiovascular diseases (4). Leukocytes contribute pro- and antiangiogenic factors but are particularly important in pathological AG (16). AG is an important component of WH, involving inflammation (46), tissue formation, and remodeling, whereby a local milieu created by the coordinated action of growth factors, cytokines, enzymes, extracellular matrix (ECM) components, and inflammatory cells interacts with the injured tissue, the result of which is the reestablishment of a physiological barrier (16).

Recent analysis of the RCMV-induced cellular transcriptome demonstrated that a significant number of RCMV-dysregulated genes are involved in WH/AG (39). We hypothesize a mechanism of CMV-accelerated TVS/CR whereby virally induced secretion of cytokines and growth factors is involved. Here, we demonstrate that the HCMV secretome contains >1,000 virally induced cellular proteins and promotes AG and WH, suggesting that HCMV accelerates TVS/CR indirectly through the production of AG and WH factors.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Preparation of the serum- and virus-free HCMV secretomes. HCMV strains AD169 and TR were produced by infected normal human dermal fibroblasts (NHDF). Infected cell supernatants were recovered at maximum cytopathic effect and cleared of cellular debris by centrifugation (10,000 x g, 15 min). Virus was pelleted through 10% sorbitol (Sigma, Inc.) in phosphate-buffered saline at 22,000 rpm (Beckman model SW28 unit) for 60 min. Pelleted virus was resuspended in serum-free Dulbecco's modified Eagle medium and titers were determined with NHDF (45). UV-inactivated HCMV was obtained by exposing 10⁸ PFU in 3 ml Dulbecco's modified Eagle medium in a 10-cm² dish for 4 min at 999 mJ/cm² in a Fisher model UVXL-1000 machine. UV-inactivated virus was tested for replication competence by NHDF infection, followed by immunostaining for IE72 protein. UV-inactivated virus not exhibiting production of IE72 was the negative control in secretome experiments. Herpes simplex virus type 1 (HSV-1) stocks were obtained from David Johnson at OHSU.

Secretomes (cell and virus-free supernatants) were generated from HCMV-, UV-inactivated HCMV-, HSV-1-, and mock-infected cells, as follows: 15 15-cm dishes of 80% confluent NHDF were washed three times with serum-free M199 medium and incubated for 12 h in serum-free EC medium (SF-SFM; Invitrogen). Subsequently, NHDF were infected with HCMV, UV-inactivated HCMV, or HSV-1 at a multiplicity of infection (MOI) of 3 in SFM or mock infected for 3 h at 37°C. Cells were washed three times with SFM and incubated for an additional 48 h. Supernatants were centrifuged (10,000 x g, 15 min), followed by two high-speed centrifugations at 25,000 rpm (Beckman model SW28 unit) for 60 min each to remove virus particles. The clarified supernatants from HCMV- and mock-infected cells were tested for infectious virus by plaque assay and immunostaining for IE72. Supernatants devoid of infectious virus and IE72 were used in secretome experiments. Additional secretomes were derived from cells treated with the HCMV DNA polymerase inhibitor foscarnet (0.5 mM) to determine whether secretome effects were mediated by a late viral gene.

Angiogenesis assay. Secretome AG activity was evaluated by using a modification of the standard Matrigel in vitro tubule formation assay (7). Primary human umbilical vein EC (HUVEC) monolayers were nutrient starved for 3 h in SF-SFM before they were resuspended and plated (10⁴ cells/well in 50 µl of SF-SFM) on preformed growth factor-reduced Matrigel plugs (BD Biosciences) in a 24-well tray. Subsequently, 300 µl of secretomes was added to triplicate wells. Control wells received either 300 µl of SF-SFM (negative control) or complete SFM (positive control). After a 24- or 48-h incubation, wells were analyzed for the formation of tubule structures by examination with a phase-contrast inverted microscope (10x objective), and the center of each well was digitally photographed. To quantitatively compare levels of tubule formation, digital images were analyzed for their degree of branching by counting branch points and for the extent of their network formation by counting enclosed polygonal spaces delimited by tubules and branch points (lumens). Data were compared for significance using Student's t test. P values of 0.05 were considered significant. Trays were incubated for 2 weeks, in some instances, to determine the long-term stability of neovessels.

WH assay. Secretome WH activity was evaluated by using an electric WH assay based on electric cell substrate impedance sensing (Applied Biophysics Inc., Troy, NY) (15). Primary HUVECs were plated in 8-well plates, in which each well contained a single active gold electrode (250-µm diameter). Slide arrays were treated with 10 mM cysteine, precoated with 1% gelatin, seeded with HUVECs (10⁵ cells/well) in complete SFM (SFM containing 10% human serum and 25 µg/ml EGCS), and incubated overnight to establish confluent monolayers. Cells were starved for 3 h in serum- and endothelial cell growth supplement-free SFM (SF-SFM) before 400 µl of secretomes was added to sets of four wells. SF-SFM and complete SFM were used as negative and positive controls, respectively. Immediately after the application of supernatants, arrays were placed in electrode holders, and confluence over the central electrode was verified by recording resistance (in ohms) to a constant current source (1 mA alternating current, 4 kHz) for 5 min. An elevated field voltage spike (2.5 V, 40 kHz for 30 s) was applied to monolayers in three of each four-well set to kill cells over each electrode. The fourth well was left unwounded to serve as a reference for an intact monolayer. Electrical wounding resulted in an immediate drop in resistance, and the subsequent rise in resistance due to migration and repopulation of the wounded area was monitored at 30-s intervals for 40 h. Data were collected by computer and imported into Excel software (Microsoft) for analysis.

Mass spectrometry analysis. HCMV- or mock-infected secretomes were collected, and volumes were reduced 10-fold by lyophilization, followed by the removal of low-molecular-weight species (<1 kDa) by using a PD-10 desalting column. Protein fractions were combined and lyophilized to further reduce volume 20-fold. Soluble proteins were denatured using 8 M urea and reduced with 5 mM tributylphosphine. Sequencing-grade trypsin was added to samples at a 1:50 (wt/wt) trypsin-to-protein ratio, samples were incubated for 5 h at 37°C, and incubation was terminated by boiling samples for 5 min.

Peptide samples were subjected to strong cation exchange chromatography using a PolyLC polysulfoethyl A 200-mm by 4.6-mm column (Columbia, MD), preceded by a 10-mm by 4.6-mm guard column with a flow rate of 1 ml/min. The separations were performed with an Agilent 1100 high-performance liquid chromatography separation system, with mobile phases consisting of solvent A (10 mM ammonium formate, 25% acetonitrile [ACN] [pH 3.0]) and solvent B (500 mM ammonium formate, 25% ACN [pH 6.8]). The injected sample was run under isocratic conditions for 10 min at 100% solvent A, followed by an initial gradient from 100% solvent A to 50% solvent B for 50 min. A steeper gradient to 100% solvent B lasting 10 min was then performed, and the sample was held isocratically at 100% solvent B for 15 min. A total of 25 fractions were collected and lyophilized to dryness.

The reversed phase capillary liquid chromatography system is composed of a 150-µm-inside-diameter by 360-µm-outside-diameter by 65-cm-long capillary (Polymicro Technologies Inc., Phoenix, AZ) fitted with a 2-µm retaining mesh and packed with 5 µm Jupiter C₁₈ stationary phase (Phenomenex, Torrence, CA). Mobile phase C consists of 0.05% trifluoroacetic acid (TFA) and 0.2% acetic acid, and mobile phase D consists of 0.1% TFA and 90% ACN. The exponential gradient mixing of mobile phase C with mobile phase D (1.8 µl/min) begins while pressure is maintained at 5,000 psi for 20 min following a 10-µl injection of the sample (1.0 µg/µl). The capillary column is interfaced with a Finnigan-LTQ ion trap mass spectrometry (MS) unit (ThermoElectron, San Jose, CA) using an electrospray ionization source. The initial MS scan utilizes an m/z range of 400 to 2,000, after which 10 of the most abundant ions were selected for tandem MS (MS-MS) analysis. Dynamic exclusion was used to prevent repeated analysis of the same high-abundance ion.

SEQUEST software was used to match the MS-MS fragmentation spectra with peptides from the most recent version of the IPI human database (8). The criteria selected for filtering are based on a human reverse database search which provides a >95% estimated level of confidence for peptide identification and higher levels for proteins identified with multiple peptides (32). Briefly, protein identifications were retained if their identified peptide met the following criteria: (i) a delta correlation (DelCN) value of 0.10 and (ii) a cross-correlation (Xcorr) score of 1.6 for charge state 1⁺ and full tryptic peptides; an Xcorr score of 2.4 for charge state 2⁺ and full tryptic peptides; an Xcorr score of 4.3 for partial tryptic peptides; an Xcorr score of 3.2 for charge state 3⁺ and full tryptic peptides; and an Xcorr score of 4.7 for partial tryptic peptides. ProteinProphet software was used to remove redundant protein identifications (26).

Human cytokine protein array. RayBio human cytokine array G Series 2000 antibody arrays (RayBiotech, Norcross, GA) were used to assay secretomes generated from control and HCMV-infected cell cultures. A total of 174 factors were evaluated (for a complete list of factors, refer to www.raybiotech.com/G_Series.asp). Triplicate biological replicates of HCMV and mock secretomes were concentrated fivefold using Amicon Ultra-4 (Millipore), and 50 µl of sample was incubated on the array. Arrays were processed according to the manufacturer's instructions, scanned using a Bioscience GeneScan Lite laser scanner, and analyzed using Imagene version 3.5.1 digital processing software (Biodiscovery). Data were subtracted for local background and normalized to internal positive and negative controls. Cutoff values for detection were set at an average intensity of 500 over that of the background. Mock- and HCMV-infected samples were compared for significance using Student's t test. P values of 0.05 were considered significant.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
HCMV induces the secretion of AG and WH factors. We have shown that RCMV accelerates the development of TVS/CR in rat heart allografts, although widespread infection is not observed with transplanted organs (39). A comparison of transcriptomes from infected and uninfected heart allografts indicated that RCMV upregulated host genes involved in AG and WH, suggesting that RCMV may accelerate TVS formation through the induction and secretion of AG/WH factors. To determine if CMV induces secretion of WH and AG factors, we analyzed the protein content of supernatants obtained from HCMV-infected cells by gel-free liquid chromatography (LCQ)-MS-MS analysis. We identified >1,200 proteins, 800 having >2 peptide hits in the HCMV-infected and mock-infected secretomes. A comparison of two separate strains of HCMV with mock-infected secretomes in three separate experiments revealed >1,000 proteins specific or highly enriched in the viral secretome, >260 proteins common to both HCMV- and mock-infected secretomes, and >225 proteins specific for the mock-infected secretome. An analysis of HCMV secretomes for viral proteins indicated only minor amounts of a few proteins (10 proteins) in which only 4 were identified with >2 peptide hits including UL32 (pp150), UL44, UL122, and UL123. The absence of viral proteins, as well as the lack of infectious virus, in the HCMV secretome indicated that supernatants contained no virus. Interestingly, pp150, the sole structural protein detected in the viral secretome, was detected by Western blotting as a 65-kDa species, suggesting that the secreted protein may be a cleavage product of the larger 150-kDa species (data not shown).

Pathway analysis indicated that many (100 proteins) of the AG/WH proteins were present in the HCMV secretome, i.e., proteins involved in transforming growth factor β (TGF-β) and other growth factor signaling pathways, cytokines, and chemokines, factors involved in ECM remodeling including a number of matrix metalloproteinases (MMP-1, -2, -3, -10, -12, and -19), cathepsins (B, D, F, K, L, and S), and tissue inhibitors of matrix metalloproteinases (TIMP-1, -2, and -3) (Table 1). These proteins act by conditioning the extracellular environment, promoting WH and AG through cellular growth and differentiation by remodeling the ECM and activating latent cellular growth factors (2, 10). A number of proteins that mediate integrin signaling pathways were also present in the HCMV secretome, such as laminins, widely distributed ECM proteins involved in cell adhesion signaling (10).

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TABLE 1. Numbers of cellular proteins identified by LC-Q-MS-MS in the HCMV secretomea

To validate and quantify MS studies, we analyzed the HCMV secretome for 174 common cytokines/growth factors, using RayBiotech human cytokine antibody arrays. In three replicated experiments, we detected 144 out of 174 proteins in both the HCMV and the mock secretomes. A comparison of the viral and mock secretomes indicated 36 significantly induced factors and 8 downregulated factors (Table 2). The most highly abundant WH/AG factors in the HCMV secretome included the cytokines/chemokines such as interleukin-5 (IL-5), IL-6, osteoprotegerin, granulocyte-macrophage-colony stimulating factor (GM-CSF), tumor necrosis factor (TNF) receptor superfamily member 18 (GITR ligand), monocyte chemoattractant protein 1 (MCP-1), MIP-1 and MIP-1β, RANTES, MCP-3, HCC-4, MIP-3, GRO, IL-8, IP-10, and ITAC (Table 3). HCMV infection also induced the release of receptors including EGF-R, TNF-R1 and -R2, ICAM-1, PECAM-1, and VE-cadherin. A number of growth factors were induced in the infected cells, including glial cell line-derived neurotrophic factor (GDNF), placental growth factor (PIGF), platelet-derived growth factor (PDGF)-AA, -AB, and -BB, and TGF-β1 and hepatocyte growth factor (HGF), as well as many ECM modifiers (MMP-1, TIMP-1, TIMP-2, TIMP-4, and uPAR) and the angiogenic RNase angiogenin. GDNF was induced 225-fold in HCMV secretome compared to that in mock secretome, which is interesting since this member of the GDNF family of neurotrophic factors is a potent promoter of cell survival. This analysis provides the first glimpse into the complexity of the HCMV secretome and reveals that HCMV infection increases the release of a number of crucial WH/AG factors.

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TABLE 2. HCMV-induced and -repressed secretome factorsa

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TABLE 3. Most abundant factors present in the HCMV secretomea

HCMV secretome induces angiogenesis in EC. In vitro angiogenesis assays rely on the fact that endothelial cells cultured on a supportive matrix such as Matrigel form vessel-like tubules in the presence of AG growth and stabilization factors. To determine if the abundance of AG factors in the HCMV, but not the mock-infected secretome, stimulated efficient angiogenesis, we employed a modification of a standard in vitro angiogenesis assay in which the HCMV or mock-infected secretome was the sole source of AG factors (7). For these experiments, low-passage primary HUVEC were nutrient starved in SFM prior to harvest and plating onto 24-well trays containing polymerized plugs of growth factor-reduced Matrigel in the presence of mock and HCMV secretomes. Controls included HUVEC grown in SF-SFM (negative control) or complete SFM (positive control). The extent of angiogenesis was quantitated by determining the number of lumens delimited by intact capillary-like tubule structures, as well as the number of interconnecting branch points. Results with control supernatants (complete SFM and SF-SFM) confirmed that exogenous angiogenic factors are required to support the formation of a robust capillary network when EC are plated on growth factor-reduced Matrigel (Fig. 1). HUVEC cultured for 24 h in complete SFM aligned to form a meshwork of anastomosing tubules with multinodal branch points and enclosed lumens. In contrast, EC cultured in SF-SFM were unable to form a consistent network of interconnecting tubules, with many cells generating incomplete tubes or aggregating in clumps. EC cultured in the presence of the HCMV secretome supported the formation of an extensive polygonal capillary network, while networks formed in the presence of the mock secretome resembled those formed in the presence of SF-SFM (Fig. 1). Vessels formed in the presence of HCMV secretome were more defined than those formed in complete SFM, suggesting the presence of a potent AG stimulus in HCMV secretome. After 48 h, capillary networks formed in SF-SFM or the mock secretome began to degenerate, with tubules retracting into aggregates (Fig. 2A and B). Networks induced by complete SFM also degenerated, as evidenced by destabilization of the initially robust tubules, with visible individual EC. In contrast, networks induced by the HCMV secretome remained stable, with smooth intercellular junctions.

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FIG. 1. HCMV supernatants induce angiogenesis at 24 h. Primary HUVEC were serum starved for 3 h in SF-SFM before they were harvested and resuspended in this medium (105 cells/ml). Cells (104 cells/well) were introduced into 24-well plates containing polymerized plugs of growth factor-reduced Matrigel in the presence of 300 µl of control and test supernatants. Test supernatants were concentrated secretomes generated from mock- and HCMV strain AD169-infected cells, using Amicon Ultra-4 filters. Control supernatants included those cultured in SF-SFM (10x concentrated) and 1x complete SFM. Each supernatant was tested in quadruplicate. (A) Quantitative measures of angiogenesis consisted of the numbers of lumens and branch points. (B) Representative examples of each culture condition are shown as a low-power image (magnification, x10). (C) High-power images emphasize differences in vessel integrity between conditions (magnification, x20). The data presented in this figure are a representation of three individual experiments.

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FIG. 2. HCMV supernatants stabilize tubule formation at 48 h and at 2 weeks. An angiogenesis assay was performed using primary HUVEC, as previously described, at 48 h postplating. (A) Low-power (magnification, x10) images show samples grown in the presence of control (SF-SFM alone or complete SFM containing human serum and ECGS) and test (mock or HCMV AD169) secretomes. (B) High-power (magnification, x20) images under complete SFM and HCMV AD169 secretome conditions are shown. (C) Tubule survival is shown after 2 weeks on Matrigel in the presence of complete SFM or HCMV AD169 secretome (magnification, x20). The data presented in this figure are a representation of three individual experiments.

To test the degree of stabilization afforded by the HCMV supernatants, trays were maintained for 2 weeks. After this time, networks formed in the presence of SF-SFM or the mock secretome had completely deteriorated (not shown), and those induced by complete SFM had degenerated severely, while those induced by the HCMV secretome remained intact (Fig. 2C). Results obtained with both the short- and the long-term AG assays indicated that HCMV secretome contains factors that promote angiogenesis and allow for the stabilization of neovessels. This activity requires HCMV replication, since the secretome of UV-inactivated HCMV-infected cells failed to support AG (data not shown).

The HCMV secretome induces wound healing in EC. The HCMV secretome contained chemokines and growth factors involved in WH events. To quantify the ability of the HCMV secretome to accelerate WH, we used automated electric cell substrate impedance-sensing technology to electrically wound cell monolayers and measure wound healing in real time (5, 15). For these assays, HUVEC were grown to confluence in 8-well chamber slides containing 250-µm-diameter gold electrodes. Figure 3A shows the appearance of unwounded HUVEC plated over an electrode in complete SFM, the effect of the voltage spike on the monolayer at the time of wounding (0 h), and the repopulation of the wound after 3 and 20 h. Figure 3B shows a typical electric cell substrate impedance-sensing (ECIS) trace for an unwounded monolayer of primary HUVEC cultured in complete SFM and a duplicate monolayer subjected to electrical wounding and subsequent wound healing. As expected, electrical wounding resulted in an immediate drop in resistance, to the level of an open electrode. In complete SFM culture, the wound begins to repopulate by 3 h postwounding and is complete in about 6 h. To specifically measure the WH capacity of mock and HCMV secretomes, primary HUVEC were plated in 8W1E arrays, in complete SFM, and incubated overnight to allow establishment of a confluent monolayer. Cells were serum starved prior to the addition of secretome samples from mock- and HCMV-infected NHDF to duplicate wells. Secretomes from NHDF infected with UV-inactivated HCMV or with live HCMV in the presence of foscarnet or from NHDF infected with HSV-1 were also tested. SF-SFM and complete SFM cultures were included as negative and positive controls, respectively. Immediately after supernatants were applied, arrays were placed in the electrode holders, and resistance was measured to confirm the existence of confluent monolayers and the absence of any nonspecific toxicity. Electrical wounding and subsequent wound healing were then monitored via continuous ECIS resistance measurements (Fig. 3B and C). Cells wounded and recovered in the presence of the HCMV secretome were able to repopulate the wound with the same kinetics as those in complete SFM, although steady-state resistance levels were slightly lower. In contrast, cells cultured in SF-SFM, mock secretome, UV-inactivated HCMV secretomes, or foscarnet-treated HCMV-infected cells were unable to repopulate the electrode, over the 20 h measured (Fig. 3B). Cells incubated with a secretome derived from HSV-1-infected cells also failed to mediate WH, suggesting that the effects are specific for HCMV (Fig. 3C). These data indicated that the HCMV secretome contains factors that effectively promote cell migration into a mechanical wound. Neither the secretomes derived from cells infected with UV-treated virions or the secretomes from HCMV-infected cells treated with foscarnet promoted WH, indicating that the ability of the HCMV secretome to mediate WH is due to active viral replication. This suggests that a late-kinetic class of HCMV gene(s) is involved in the generation of secretome WH factors; this correlates with studies of human transplant patients, as well as our observations for rat heart transplants in which ganciclovir prolongs graft survival of CMV-infected recipients.

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FIG. 3. Wound healing activity of the HCMV secretome. EC were grown to confluence on 8W1E ECIS arrays and exposed to test supernatants before they were electrically wounded. (A) Low-magnification images of ECIS electrodes containing unwounded and wounded EC grown in the presence of complete SFM at the time of wounding (0 h) and at 3 and 20 h postwounding. (B) Wound healing, as indicated by increasing resistance, is plotted as a function of time. Healing traces for duplicate wells are shown for the mock secretome (Mock, green trace) and HCMV strain AD169 secretome (HCMV, red trace). Also shown is the wound healing activity of secretomes from HF infected with UV-inactivated HCMV (UV-HCMV, pink trace), or infected (HCMV/Foscarnet, orange trace) or mock-infected (Mock/Foscarnet, light blue trace) in the presence of foscarnet. Control traces include a negative control (SF-SFM, yellow), a positive control (Complete SFM, dark blue), and a constant resistance trace measured from a confluent unwounded monolayer (Unwounded, black). Cells exposed to the HCMV secretome show wound repopulation within 8 to 10 h, whereas cells exposed to the mock secretome repopulate the wound inefficiently. Secretomes from UV-inactivated HCMV or HCMV plus foscarnet similarly fail to repopulate the wound, indicating that infectious virus and late gene expression are required for the production of wound healing factors. (C) Wound healing in response to the mock secretome (Mock, green), the HCMV AD169 secretome (HCMV, red), and the HSV-1 secretome (HSV-1, yellow). Also shown is the wound healing activity of secretomes from HF infected with UV-inactivated HCMV AD169 (UV-HCMV, light blue) and negative control (SF-SFM, pink), a positive control (Complete SFM, dark blue), and a constant resistance trace measured from a confluent unwounded monolayer (Unwounded, black). Cells exposed to the HCMV secretome show wound repair within 6 to 10 h, whereas cells exposed to the HSV-1 or mock secretomes repopulate the wound inefficiently, indicating that the production of wound healing factors is specific for HCMV infections. The data presented in this figure are a representation of three individual experiments.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
In this report, we used in vitro assays to show that virus- and cell-free supernatants from HCMV-infected cells induce the secretion of AG/WH-promoting factors. Interestingly, the HCMV secretome also contains factors that promote long-term EC survival. This occurs to a far greater extent in cells cultured in the presence of the HCMV secretome than in cells cultured in serum. The generation of the HCMV secretome requires active viral replication and is sensitive to ganciclovir. This parallels observations for human heart transplant recipients, where ganciclovir prolongs graft survival in the presence of CMV infection (44). Together, these findings demonstrate that CMV infection induces AG/WH via a paracrine effect.

Vascular disease is a complex process that initially intended to heal an injured blood vessel but subsequently went awry. Vascular disease involves multiple components including the primary injury event, the local response to injury through platelet aggregation, and the production of cytokines and growth factors, followed rapidly by inflammation. The next step is SMC migration and proliferation, which can ultimately lead to vessel occlusion. HCMV infection of the vasculature may contribute to any one or all of these components, thereby accelerating the disease process. Since HCMV infects many of the cell types involved in vascular disease, the initial vessel injury may result from the direct effects of immune clearance of infected cells and/or the expression of pathogenic viral proteins. Our data suggest that indirect mechanisms, mediated by HCMV, occur through the release of growth/differentiation factors that tip the balance from a natural healing process to a pathological one. Here, we show that HCMV-infected cells secrete a number of cytokines (IL-1 and IL-1β, IL-5, IL-6, GM-CSF, osteoprotegerin, TNF-, and TNF-R1 and -R2), and chemokines (MCP-1, -3, and -4; MIP-1 and MIP-1β, RANTES, IL-8, IP-10, ITAC, SDF, GRO-, MIP-3, and midkine). Importantly, CMV carries chemokine homologues that also recruit and stimulate a multitude of host cellular infiltrates that mediate the inflammatory response (27, 36). RCMV-infected cardiac allograft tissues contain increased numbers of cellular infiltrates including MØ and CD4/CD8 T cells, and the presence of these cells parallels the upregulation of chemokines and cytokines (38). Factors constituting the HCMV secretome are very similar to those for the genes that were upregulated during the critical phase of RCMV-accelerated TVS in our heart transplant model (39). The production of chemokines and cytokines by infected cells likely exacerbates the initial response to vessel injury and enhances the inflammatory process.

Increasing evidence supports a role for AG and WH in a number of vascular diseases. For example, intraplaque neovascularization is a common feature of atherosclerosis, and many proangiogenic factors are found in human atherosclerotic lesions (17). Increased adventitial neovascularization is also seen at sites of neointimal hyperplasia following arterial or venous stenting (34). Inflammatory cells and SMC within lesions are important local sources of proangiogenic factors (21). Not surprisingly, antifibrotic/antiangiogenic agents like rapamycin decrease neointimal hyperplasia in coronary artery stents (31, 40, 41). The pathological processes involved in vascular disease parallel many of the cellular/molecular events that mediate normal WH/AG. WH is a complex sequential process involving inflammation, tissue formation, and remodeling and resulting in the reestablishment of an anatomical or physiological barrier (16). The process requires the influx of inflammatory cells and the coordinated actions of growth factors, cytokines, enzymes, and ECM components interacting with the injured tissue. Given the roles of inflammation, new vessel growth, and tissue remodeling in WH, it is not surprising that many of the same stimulatory/inhibitory factors promote both AG and WH (46).

The HCMV secretome contains a number of factors that contribute to WH and AG, including growth factors (angiopoietin, angiotensinogen, FGF, GDNF, HGF, IGF-BP, osteopontin, PDGF, PIGF, SPARC, TGF-β, thrombospondin, and VEGF), ECM and ECM-modifying proteins (collagen, fibronectin, GDF-15, laminin, MMPs, TIMPs, thrombospondin, and uPAR), the ELR containing CXC chemokines (IL-8 and GRO-), cytokines (IL-1 and IL-1β, IL-5, IL-6, GM-CSF, osteoprotegerin, TNF-, and TNF-R1 and -R2), enzymes (cathepsin B, D, F, K, L and S, PAI-1, PAI-2, and SERPIN 12), and adhesion molecules (ALCAM, ICAM, PECAM, and E-cadherin). Many of these regulatory factors promote EC migration and proliferation. TGF-β has profound effects on developmental processes, including cell proliferation, differentiation, adhesion, and inflammatory responses, as well as on WH. TGF-β acts on nontransformed tumor cells to suppress antitumor immune responses and augment AG. VEGF is also a potent activator of EC proliferation, and the small RNase angiogenin is important for EC invasion. In addition, the extracellular endopeptidases cathepsin B, K, and L may directly contribute to angiogenesis and recruitment of EC progenitors during AG and WH. Many AG/WH factors present in the HCMV secretome regulate these processes to both a positive and a negative degree, a finding corroborated by our demonstration that the HCMV secretome not only promotes EC tubule formation but also stabilizes the tubule network for 2 weeks. The positive control for these AG assays, cells treated with serum, promoted efficient tubule formation but failed to sustain a healthy existence on a long-term basis, suggesting that the HCMV secretome contains both formation and stabilization factors. Many components of the HCMV secretome contain both of these properties; for example, the ELR containing CXC chemokines IL-8 and GRO, which promote angiogenesis, and the non-ELR containing the CXC chemokine IP-10, which inhibits angiogenesis but may promote EC stability. A number of other HCMV secretome factors such as thrombospondin-1 and TIMPs inhibit angiogenesis, eliciting vessel-stabilizing effects. A number of the constituents of the HCMV secretome were also present in RCMV-infected heart allografts, including growth factors (angiopoietin, FGF, GDNF, HGF, IGF-BP, osteopontin, PDGF, PIGF, SPARC, TGF-β, thrombospondin, and VEGF), ECM modifiers (cathepsins, fibronectin, MMPs, PAI-2, TIMPs, and uPAR), and adhesion molecules (ALCAM and ICAM-1) (39). Our combined in vitro and in vivo findings support the notion that factors secreted by CMV-infected cells tip the balance to a profibrotic state, causing vessel neointimal hyperplasia and vessel narrowing.

SMC proliferation and migration into the neointimal space are the crux of vascular lesion formation, suggesting that CMV targets these events promoting SMC accumulation. Here, we report that the HCMV secretome not only promotes AG but also has a very dramatic effect on long-term EC tubule survival. Whether the effect of the secretome is mediated directly by viral proteins or indirectly through the induction of cellular factors remains unclear. One possibility for the enhancement of EC survival occurs through a reduction in apoptosis. However, the HCMV-encoded inhibitors of cell death, vICA and vMIA, were not detected in our HCMV secretome proteomics study. In addition, it is unclear whether vMIA and vICA can enter cells or whether they must be produced within target cells. SMC accumulation may occur through increased migration and proliferation at the site of vascular injury. Previously, others have shown that HCMV infection of EC increases the expression of FGF and PDGF-BB, both of which are potent stimuli in SMC proliferation (13, 14, 20, 33). The HCMV secretome contains additional SMC growth factors that stimulate proliferation, including angiopoietin, PlGF, GDNF, FGF-7, PDGF, TGF-β1, and HGF. In addition, the HCMV secretome contains many proteins involved in ECM remodeling (MMPs and TIMPs) and proteins involved in the release of latent growth factors, such as uPAR and the angiogenic RNase angiogenin, which mediate SMC migration and/or proliferation. MMPs remove the basement membrane around vascular SMC and facilitate contacts with the interstitial matrix. This process promotes a phenotypic change from quiescent, contractile vascular SMC to cells capable of migrating and proliferating to mediate repair.

Clinical studies of transplant and angioplasty patients have linked HCMV to the acceleration of vascular disease; however, the mechanism(s) is not understood. The lack of feasible human models is the primary difficulty in determining the mechanisms of HCMV infection-accelerated vascular disease. We have developed a rat model of heart transplantation in which RCMV infection accelerates the time to development and the severity of TVS between days 21 and 28 posttransplantation (29, 38). RCMV infection of allograft recipients increases immune cell infiltration, chemokine production, and upregulation of WH and AG genes during this critical phase (39). Thus, the findings presented in this paper together with the observations for the RCMV allograft model provide a possible mechanism of CMV infection-accelerated TVS through the induction of a pathological tissue-remodeling process, which in the setting of inflammation and alloreactivity, leads to neointimal hyperplasia through the upregulation of cellular WH and AG genes.

Our understanding of the role of HCMV in accelerating vascular diseases continues to develop. Progress has been made through extensive epidemiology, the use of animal models, and the use of in vitro models that dissect processes related to the clinical scenario. While precise mechanisms have yet to be fully elucidated, it has now become clear that HCMV modifies the extracellular host environment through the production and release of biologically active cellular factors including growth factors, cytokines, and ECM-modifying enzymes. We demonstrate here that the HCMV secretome is capable of mediating angiogenesis and wound healing, which are important processes driving vascular disease formation. Many host factors identified in the HCMV secretome are also upregulated by RCMV infection in rat allograft hearts, suggesting that our in vitro findings parallel the in vivo setting. Future studies to identify and target specific viral genes mediating production of the HCMV secretome will be vital for the possible prevention or abrogation of HCMV-associated vasculopathy.

 
S. L. Orloff was supported by grants from the Department of Veterans Affairs and from the National Institutes of Health (HL 66238-01). NIH grants were also awarded to D. N. Streblow (HL083194), A. V. Moses (RR00163), and J. A. Nelson (AI21640, HL65754, and HL71695). D. N. Streblow is also supported by an AHA Scientist Development grant.

J.D. performed research and analyzed data; D.N.S. analyzed data and wrote the manuscript; A.V.M. performed experiments and wrote the manuscript; J.M.J. performed research and analyzed data; C.N.K. performed research; D.C. analyzed data; R.D.S. analyzed data; S.L.O. wrote the manuscript; and J.A.N. wrote the manuscript.

 
* Corresponding author. Mailing address: Oregon Health Sciences University, Vaccine and Gene Therapy Institute, Mail Code VGTI, 505 N.W. 185th Ave., Beaverton, OR 97006-3499. Phone: (503) 494-7769. Fax: (503) 494-2441. E-mail: nelsonj{at}ohsu.edu

Published ahead of print on 30 April 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

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Journal of Virology, July 2008, p. 6524-6535, Vol. 82, No. 13
0022-538X/08/$08.00+0     doi:10.1128/JVI.00502-08
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