Human cytomegalovirus (HCMV) is a prevalent betaherpesvirus that is asymptomatic in healthy individuals but can cause serious disease in immunocompromised patients. HCMV is also the leading cause of virus-mediated birth defects. Many of these defects manifest within the central nervous system and include microcephaly, sensorineural hearing loss, and cognitive developmental delays. Nitric oxide is a critical effector molecule produced as a component of the innate immune response during infection. Congenitally infected fetal brains show regions of brain damage, including necrotic foci with infiltrating macrophages and microglia, cell types that produce nitric oxide during infection. Using a 3-dimensional cortical organoid model, we demonstrate that nitric oxide inhibits HCMV spread and simultaneously disrupts neural rosette structures, resulting in tissue disorganization. Nitric oxide also attenuates HCMV replication in 2-dimensional cultures of neural progenitor cells (NPCs), a prominent cell type in cortical organoids that differentiate into neurons and glial cells. The multipotency factor SOX2 was decreased during nitric oxide exposure, suggesting that early neural differentiation is affected. Nitric oxide also reduced maximal mitochondrial respiration in both uninfected and infected NPCs. We determined that this reduction likely influences neural differentiation, as neurons (Tuj1+ GFAP− Nestin−) and glial populations (Tuj1− GFAP+ Nestin−) were reduced following differentiation. Our studies indicate a prominent, immunopathogenic role of nitric oxide in promoting developmental defects within the brain despite its antiviral activity during congenital HCMV infection.
IMPORTANCE Human cytomegalovirus (HCMV) is the leading cause of virus-mediated congenital birth defects. Congenitally infected infants can have a variety of symptoms manifesting within the central nervous system. The use of 3-dimensional (3-D) cortical organoids to model infection of the fetal brain has advanced the current understanding of development and allowed broader investigation of the mechanisms behind disease. However, the impact of the innate immune molecule nitric oxide during HCMV infection has not been explored in neural cells or cortical 3-D models. Here, we investigated the effect of nitric oxide on cortical development during HCMV infection. We demonstrate that nitric oxide plays an antiviral role during infection yet results in disorganized cortical tissue. Nitric oxide contributes to differentiation defects of neuron and glial cells from neural progenitor cells despite inhibiting viral replication. Our results indicate that immunopathogenic consequences of nitric oxide during congenital infection promote developmental defects that undermine its antiviral activity.
The prevalent betaherpesvirus human cytomegalovirus (HCMV) can cause serious disease during infection. Individuals at higher risk of disease are those that are immunocompromised or receiving immunosuppressive therapies (1). Additionally, congenital HCMV infection occurs in 0.5 to 1% of all live births and is the leading cause of virus-mediated birth defects as reported for the United States and United Kingdom (2). Congenital infection occurs through primary or secondary infection of a pregnant person and subsequent vertical transmission to the developing fetus. A subset of these infants will exhibit neurological manifestations, either at birth or later in life, that include microcephaly, seizures, sensorineural hearing loss, and other long-term developmental defects (3–5). The mechanisms behind this pathogenesis are not well defined but involve infection of the central nervous system (CNS) (6–8).
HCMV has wide cell tropism and replicates in various cell types, including fibroblasts, epithelial cells, endothelial cells, trophoblasts, and cells within the CNS, such as neural progenitor cells (NPCs) (9). NPCs are primarily found in the subventricular zone of the brain and differentiate into the main cell types of the CNS, including neuron and glial cells (10). NPCs derived from embryonic and induced pluripotent stem cells (iPSCs) are susceptible to HCMV infection; however, the stage of differentiation impacts susceptibility, with less differentiated cells being more susceptible (11, 12). Infection decreases proliferation and results in abnormal differentiation of NPCs (13–16). Viral protein expression has been demonstrated to contribute to abnormal differentiation both directly and indirectly. HCMV IE1 sequestering of STAT3 to the nucleus results in decreased multipotency factor SOX2 expression (17–19). Liu et al. (14) demonstrated IE1 promotes ubiquitination and proteasomal degradation of the transcriptional regulator Hes1.
Identifying the mechanisms behind HCMV pathogenesis during fetal development was largely confined to 2-dimensional cultures, due to species specificity, until the recent development of iPSC-derived cortical organoids. This 3-dimensional tissue model recapitulates many structural components and the transcriptional profile of the developing fetal brain (20). This tissue has recently been used to model CNS defects during congenital HCMV infection. We previously demonstrated that cortical organoids infected with HCMV have altered neural layering and disrupted calcium signaling (21). Brown et al. (22) showed that HCMV-infected iPSCs retain the ability to differentiate into cortical organoids but exhibited gross morphological differences compared to mock-infected iPSC-derived organoids. Additionally, expression of genes involved in neurogenesis and brain development are substantially dysregulated during HCMV infection of cortical organoids (23). The use of cortical organoids to model HCMV infection of the developing brain allows more complex investigation into contributing factors during infection such as the innate immune system.
During HCMV infection, innate immune cells produce several nonspecific antiviral molecules, including nitric oxide via the enzyme nitric oxide synthase 2 (NOS2) (24, 25). NOS2 produces picomolar to micromolar levels of nitric oxide (26, 27), thereby exposing both infected and uninfected cells to various concentrations of nitric oxide. High levels of nitric oxide can inhibit enzymes containing iron-sulfur clusters, including complex I and complex IV of the electron transport chain (ETC) and aconitase, an enzyme in the tricarboxylic acid (TCA) cycle (28–30). These inhibitions impact several aspects of cellular function, such as proliferation, differentiation, metabolism, mitochondrial function, and energy production. In plated NPCs from mice, nitric oxide can enhance or inhibit proliferation in a concentration-dependent manner (31, 32). Further, Bergsland et al. (33) demonstrated that neuron differentiation was decreased following nitric oxide exposure. However, the full impact of nitric oxide on events within the developing brain is largely undefined.
There is evidence that nitric oxide controls viral infections within the brain while also contributing to disease. Nitric oxide inhibits many RNA and DNA viruses (34), including herpesviruses such as herpes simplex virus-1 (HSV-1), herpes simplex virus-2 (HSV-2), and Epstein-Barr virus (EBV) (35–38). Despite this antiviral activity, nitric oxide promotes immunopathogenesis during infection. During HSV-1 infection, exposure to a nitric oxide donor contributed to amyloid beta (Aβ) accumulation in both neuronal cultures and mouse brains (39). In mice with West Nile virus-induced encephalitis, inhibition of nitric oxide production with a selective NOS2 inhibitor increased survival (40). Wang et al. demonstrated that the conditioned media from Zika virus-infected microglia, which contains nitric oxide, decreased neuronal differentiation (41).
Nitric oxide is also inhibitory to HCMV replication and plays an important role in regulating infection (42–46). A case study described an individual with NOS2 deficiency who succumbed to HCMV infection despite evidence of past pathogenic infections that did not prove fatal (47). We have recently demonstrated that nitric oxide inhibits HCMV replication in human fibroblasts and retinal pigment epithelial cells through a multifactorial mechanism involving inhibition of mitochondrial respiration and dysregulation of metabolism (48). It is unknown how these mechanisms impact infection within the fetal brain. Congenitally infected fetal brains have necrotic foci with infiltrating macrophages and microglia that are capable of producing nitric oxide (7, 24, 25). In murine models of congenital CMV infection, embryonic mice and pups also have infiltrating NOS2-expressing macrophages within the brain (49, 50).
Nitric oxide is a potent antiviral yet is considered a double-edged sword due to its association with tissue damage. Its impact on CNS development during HCMV infection is unknown. Here, we determined the effect of nitric oxide on HCMV infection of cortical organoids and NPCs as well as immunopathogenic consequences on uninfected cells. We demonstrate that nitric oxide attenuates viral spread in cortical organoids and NPCs while dysregulating neural differentiation. Nitric oxide decreased levels of SOX2 and compromised mitochondrial function, both important regulators of neural differentiation. We demonstrate the role of nitric oxide in controlling HCMV infection while causing immunopathogenic effects that are detrimental to the developing fetal brain. These discoveries provide novel insights into neuropathogenesis observed during congenital HCMV infection.
Nitric oxide reduces HCMV spread in cortical organoids.
Modeling defects in the CNS during congenital HCMV infection have been largely limited to 2-dimensional cell culture systems due to species specificity. However, the emergence of 3-dimensional cortical organoids has allowed investigation of congenital infection in a tissue model with features of fetal forebrain (21–23). Brains from congenitally infected fetuses have necrotic regions that contain infiltrating macrophages and microglia (7). These immune cells produce nitric oxide via NOS2 in response to pathogenic infection (24–26), suggesting that nitric oxide is present in the fetal brain during congenital infection. However, the impact of nitric oxide on HCMV infection and tissue within the fetal brain is unknown. To determine the role of nitric oxide during HCMV infection in developing neural tissue, we began our studies by defining the effect of HCMV and nitric oxide on cortical organoids. We and others demonstrated previously that nitric oxide inhibits HCMV replication in cultured human fibroblasts and epithelial cells using the spontaneous-release nitric oxide donor diethylenetriamine NONOate (DETA/NO) to mimic nitric oxide production by NOS2 (48, 51). Since macrophages and microglia are not typically present in cortical organoids, we used DETA/NO to determine the impact of nitric oxide on this tissue. Cortical organoids were differentiated from an iPSC line derived from a heathy individual (52, 53). Organoids were cultured to day 35 of development, at which time they have developed defined brain region identities (54). Organoids were mock infected or infected using HCMV strain TB40/E encoding enhanced green fluorescent protein (TB40/E-eGFP) at a multiplicity of infection (MOI) of 500 infectious units (IU) per microgram of tissue. Viral stocks were produced using MRC-5 human fibroblasts (TB40/EFb-eGFP). Organoids were treated at 2 h postinfection (hpi) and every 24 h with 400 μM DETA/NO or vehicle. Organoids were treated with DETA (spent donor) as an additional control. DETA (spent donor) was prepared by incubating medium containing 400 μM DETA/NO at 37°C for 72 h (48). This incubation releases nitric oxide from DETA/NO and leaves only the parental backbone and nitric oxide oxidation products, which serve to distinguish effects of nitric oxide from these variables. GFP expression was not observed in mock-infected organoids at 11 days postinfection (dpi) (Fig. 1A). We observed an increase in GFP fluorescence and spread from 4 dpi to 11 dpi in vehicle and DETA-treated, HCMV-infected organoids (Fig. 1A), indicating efficient HCMV replication. In DETA/NO-treated, HCMV-infected organoids, GFP was dramatically decreased compared to vehicle and DETA (Fig. 1A). We quantified the mean fluorescent intensity (MFI) normalized to the cross-sectional surface area and observed an average 79% decrease during DETA/NO exposure compared to vehicle (Fig. 1B). DETA/NO-treated organoids maintained a size similar to that seen under vehicle and DETA conditions, suggesting that nitric oxide did not impact organoid growth (Fig. 1B). These data demonstrate that nitric oxide, but not DETA backbone or nitric oxide oxidation products, reduces HCMV spread in cortical organoids.
Macrophages and microglia produce nanomolar to micromolar levels of nitric oxide in response to infection (26, 27). We estimated steady-state nitric oxide levels to ensure that concentrations were within physiological conditions. We modeled steady-state nitric oxide release over 24 h from 200 and 400 μM DETA/NO using COPASI software (Fig. 1C) (55). The maximum steady-state nitric oxide concentrations from 200 and 400 μM were predicted to be 1.3 μM and 1.8 μM, respectively (Fig. 1C). These results demonstrate that cortical organoid exposure to concentrations of nitric oxide observed during infection is sufficient to reduce HCMV spread in cortical organoids.
Neural rosette structures and tissue organization are disrupted in nitric oxide-exposed cortical organoids.
Similar to the human brain, cortical organoids develop 3-dimensional structure with distinct multicellular layer identities (54). HCMV infection of cortical organoids increases cell death and disrupts tissue morphology and organization, specifically neural rosette structures (21–23). To determine if nitric oxide inhibition of HCMV spread could alleviate this disruption, we evaluated cell viability and neural rosette structure. Consistent with our previous studies (21), we observed that GFP fluorescence was largely confined to the periphery of the organoid, with limited spread to the inner layers (Fig. 2A). We first labeled for fragmented DNA using terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) in cryosectioned organoids because nitric oxide is known to induce DNA damage (Fig. 2A). We quantified TUNEL relative to Hoechst total DNA signal (Fig. 2B). In uninfected organoids, the TUNEL/Hoechst ratio was significantly increased by 2.3-fold in DETA/NO-treated organoids compared to vehicle, which did not occur in control DETA (spent donor) conditions (Fig. 2B). We observed an increased ratio of TUNEL/Hoechst across all conditions in HCMV-infected organoids similar to that seen in mock DETA/NO-treated samples (Fig. 2B). We cannot rule out the possibility that the upper limit of detection was reached for this assay. We next measured cell viability. Organoids were infected and treated as described above. At 11 dpi, organoids were dissociated to obtain single cells and viability was quantified by trypan blue exclusion. In contrast to TUNEL, mock-infected organoids had no difference in cell viability between conditions (Fig. 1C). All HCMV-infected organoids had decreased cell viability compared to mock-infected ones (Fig. 2C). The addition of DETA/NO further reduced viability, yet these differences were not statistically significant. Collectively, our data suggest that nitric oxide does increase DNA damage in uninfected samples with no detectible impact on viability. In contrast, infection increased TUNEL regardless of nitric oxide exposure as well as reduced overall viability.
Tissue sections also revealed substantial changes to organoid organization, including regions lacking TUNEL and Hoechst (Fig. 2A). The areas were present in all organoid conditions but were more evident in mock and HCMV-infected organoids exposed to nitric oxide (Fig. 2A, arrows). As the regions appeared similar to radial neural rosette structures formed by SOX2-expressing neural progenitor cells (NPCs), we labeled for SOX2, a transcription factor necessary for progenitor maintenance and a marker for NPCs (56–59). We observed that mock-infected, vehicle-treated organoids displayed SOX2-expressing cells organized in layered radial neural rosette morphology (Fig. 3A and B, arrowheads). As previously described (21), rosette structures were disrupted in HCMV-infected, vehicle-treated organoids (Fig. 3A and B, arrows). SOX2-expressing cells also formed rosette-like structures in nitric oxide-exposed cortical organoids. However, the structures lacked the morphology and layering of a typical rosette, and the overall organization of the structures was compromised (Fig. 3B, arrowheads versus arrows). DETA (spent donor) alone in mock- and HCMV-infected organoids exhibited an intermediate phenotype, with some sections displaying more typical rosette morphology and others showing disrupted rosette-like structures (Fig. 3B, arrowheads versus arrows). Our data indicate that nitric oxide, and possibly its oxidation products, disrupts cortical organoid organization. Overall, these results suggest that nitric oxide inhibits viral spread in cortical organoids yet, regardless of infection, disrupts structure and organization.
Nitric oxide attenuates HCMV replication and alters markers of NPC proliferation.
SOX2-expressing NPC rosettes were disrupted in nitric oxide exposed organoids (Fig. 3A and B). Therefore, we explored the impact of nitric oxide on NPC development and function during infection. NPCs are susceptible to HCMV infection and are significantly disrupted in HCMV-infected organoids (21, 22). We used 2-dimensional cultures of NPCs differentiated from the same healthy iPSC line from which the cortical organoids were derived. NPCs were cultured as neurospheres before dissociation and plating (60).
Nitric oxide can have cytotoxic and cytostatic effects based on concentration. Therefore, we assessed these effects in uninfected NPCs. Cells were plated subconfluently at 5 × 105 cells/well and treated with a range of 100 to 400 μM DETA/NO or vehicle or not treated. Cell viability was quantified again using trypan blue exclusion at 96 h posttreatment. Cell viability at 100 and 200 μM concentrations were similar to vehicle and no treatment conditions (Fig. 4A). However, viability of cells in monolayer cultures decreased at higher concentrations. We observed an initial increase in live-cell number in untreated and vehicle-treated NPCs; however, this increase did not occur in 100 and 200 μM DETA/NO, suggesting that nitric oxide induces cytostasis at specific concentrations. We observed a decrease in live-cell count at 300 and 400 μM concentrations (Fig. 4A).
We next examined the impact of nitric oxide on HCMV replication. To increase infection efficiency, we used TB40/E-eGFP produced in ARPE-19 epithelial cells (TB40/EEpi-eGFP) (61). NPCs were plated, incubated for 3 days, and infected with TB40/EEpi-eGFP at an MOI of 3 IU/cell. At 2 hpi, cultures were treated with 200 μM DETA/NO, DETA (spent donor), or vehicle control, which were replaced every 24 h. As previously described, the maximum steady-state nitric oxide concentration from 200 μM DETA/NO was estimated to be 1.3 μM (Fig. 1C). Viral DNA levels were quantified at 2, 48, and 96 hpi (Fig. 4B). Titers of cell-free virus from 96 hpi were determined on ARPE-19 epithelial cells (Fig. 4B). Viral DNA levels increased by 0.8 log from 2 to 96 hpi, indicating productive viral replication. Nitric oxide reduced viral DNA levels by 0.6 log and viral titers by 0.9 log at 96 hpi (Fig. 4B). TB40/EEpi-eGFP is known to be more highly cell associated. Therefore, we quantified titers of both cell-associated and cell-free virus from 96 hpi (Fig. 4C). Cell-free titers were decreased by ~0.9 log compared to cell-associated titers in both vehicle and DETA (spent donor) control. Nitric oxide reduced cell-associated and cell-free titers by 1 log and 0.8 log, respectively; however, this difference was not statistically significant (Fig. 4C). To further elucidate when nitric oxide impacts viral replication, we determined levels of viral proteins using Western blot analysis. Nitric oxide did not impact IE1 (immediate early) levels (Fig. 4D and E). However, IE2 (immediate early), UL44 (early), and pp28 (late) levels were decreased after 48 hpi during nitric oxide exposure (Fig. 4D and E). Together these data suggest that nitric oxide attenuates HCMV replication in NPCs after the onset of viral DNA synthesis, and this result is consistent with our previous studies in human fibroblasts and epithelial cells (48).
Nitric oxide is cytostatic or cytotoxic to NPCs in a concentration-dependent manner (Fig. 4A), which can have consequences for cellular differentiation (62, 63). To investigate nitric oxide-induced cytostasis, we measured steady-state levels of the cell cycle regulators cyclin-dependent kinase inhibitor p21CIP1/WAF1, which induces cell cycle arrest, and cyclin B, which promotes cell cycle progression (Fig. 5A). Plated NPCs were infected as described above at an MOI of 3 IU/cell or mock-infected and treated at 2 hpi with 200 μM DETA/NO or vehicle control. In uninfected NPCs, p21CIP1/WAF1 levels were increased during nitric oxide exposure by 3.4-, 4.1-, 3.9-, and 3.6-fold at 24, 48, 72, and 96 hpi, respectively, relative to vehicle (Fig. 5A and B). In contrast, levels remained unchanged in HCMV-infected cells (Fig. 5A and B). No changes were observed for cyclin B levels (Fig. 5A and B). These data suggest that nitric oxide exposure induces p21CIP1/WAF1 expression, which likely contributes to cytostasis of uninfected NPCs. Expression of p21CIP1/WAF1 and cell cycle exit are tightly linked with the onset of cellular differentiation (62, 63). We hypothesized that nitric oxide-induced cytostasis may interfere with NPC maintenance and differentiation in mock and HCMV-infected cultures. SOX2 is a master regulator required for maintaining multipotency and progenitor identity of NPCs (56–59, 64). Downregulation of SOX2 is associated with cell cycle exit, decreased proliferation, and terminal differentiation (59, 65). Studies have demonstrated that SOX2 levels are reduced in an IE1-dependent manner during HCMV infection of embryonic-derived NPCs (13, 19). We initiated our studies by investigating SOX2 levels during nitric oxide exposure. Plated NPCs were mock or HCMV infected as described above at an MOI of 3 IU/cell and treated at 2 hpi with 200 μM DETA/NO or vehicle control, and SOX2 levels were determined at 96 hpi by Western blot analysis (Fig. 5C). As we were interested in the separate impact of nitric oxide on uninfected and infected cultures, we set DETA/NO treatment relative to vehicle for each condition. SOX2 levels were decreased by 0.3-fold relative to vehicle in uninfected NPCs. This reduction was more dramatic in HCMV-infected NPCs, with a 0.6-fold decrease during nitric oxide exposure. Our data demonstrate that nitric oxide exposure reduces SOX2 levels and cellular proliferation and increases p21CIP1/WAF1 levels. Taken together, these results suggest that nitric oxide disrupts regulators of multipotency and differentiation.
Mitochondrial function is compromised during nitric oxide exposure.
During differentiation, NPCs undergo a metabolic shift from mainly glycolytic to oxidative phosphorylation involving mitochondrial respiration (66, 67). Nitric oxide can disrupt mitochondrial respiration directly by inhibiting complex I and IV of the electron transport chain or indirectly through aconitase inhibition (28–30). To determine if nitric oxide decreases respiration, we examined mitochondrial function using an extracellular flux assay (Fig. 6). NPCs were plated, incubated for 3 days, infected at an MOI of 3 IU/cell or mock infected, and treated at 2 hpi with 200 μM DETA/NO, DETA (spent donor), or vehicle control. At 24 hpi, a mitochondrial stress assay was performed. Briefly, basal oxygen consumption rate (OCR) of cells (Fig. 6A) is measured before sequential injections of oligomycin (ATP synthase inhibitor), carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP; uncoupler of mitochondria), and rotenone/antimycin A (complex I and III inhibitors, respectively). This assay measures basal (green), ATP-linked (blue), and maximal (yellow) OCR (Fig. 6A). The difference between basal and maximal OCR is the spare capacity of the cells. A representative plot of the mitochondrion stress assay is shown in Fig. 6B, with quantification of OCR shown in Fig. 6C to F.
Basal OCR of HCMV-infected cells was significantly increased by 79% compared to uninfected NPCs (Fig. 6C), indicating that infection increases mitochondrial respiration by 24 hpi in NPCs. However, nitric oxide exposure reduced basal OCR by 33% in infected NPCs but not in uninfected cells. ATP-linked OCR also increased by 84% in vehicle-treated, HCMV-infected cells compared to vehicle-treated, uninfected NPCs (Fig. 6D). This rate was reduced by 31% upon nitric oxide exposure of HCMV-infected cells. No differences were observed in mock-infected conditions. Maximal respiration of NPCs was increased by 39% during infection (Fig. 6E), indicating an increase in the maximum range of the cells’ response to energy demands. However, in contrast to basal and ATP-linked OCR, maximal respiration was reduced in both uninfected and HCMV-infected NPCs by a respective 55% and 54% during exposure to nitric oxide. Likewise, the spare respiratory capacity of the cells was reduced by a respective 89% and 76% in mock and HCMV-infected conditions during nitric oxide exposure (Fig. 6F). OCRs of the DETA (spent donor) did not deviate from those of vehicle for the uninfected or HCMV-infected groups and were not included in the statistical analysis (Fig. 6C to F). These data demonstrate that nitric oxide significantly suppresses HCMV-mediated increase of basal and ATP-linked respiration. Furthermore, and regardless of infection, nitric oxide reduces the capacity of NPCs to respond to energy demands through mitochondrial respiration regardless of infection.
Nitric oxide limits neuron and glial lineages during NPC differentiation.
Reduced mitochondrial respiration, decreased SOX2 expression, and cytostasis led us to hypothesize that nitric oxide restricts neuron and glial cell differentiation from NPCs. To test this hypothesis, we investigated the effect of nitric oxide with and without infection on neural differentiation at 14 days post differentiation. Cells were cultured in the absence of growth factors to allow spontaneous differentiation (16, 60). We treated them with 200 μM DETA/NO every 48 h to reduce the chance of cytotoxicity due to prolonged exposure and reduced the MOI to avoid loss of cultures due to lytic replication during the 14 days. NPCs were plated for 3 days and infected at an MOI of 0.05 IU/cell or mock infected. Subsequently, cells were treated at 2 hpi and every 48 h with DETA/NO, DETA (spent donor), or vehicle control up to 11 dpi. We assessed markers of neural populations using immunofluorescence (Fig. 7A). We used the marker Nestin to identify the progenitor population because SOX2 is reduced during HCMV infection of embryonic stem cell-derived NPCs (13). Further, we identified neuron populations by labeling for Tuj1, a class III beta-tubulin specific to neurons, and glial populations by labeling for glial fibrillary acidic protein (GFAP), a type III intermediate filament protein expressed by glial cells (Fig. 7A). In mock-infected vehicle-treated cultures, we observed expression of each developmental marker, indicating a mix of neural populations. We observed Nestin-positive cells contained small processes with large cell bodies (Fig. 7A), while Tuj1-positive cells had long, netted neurites with small cell bodies (Fig. 7A). GFAP-positive cells had long projections with a large cell body (Fig. 7A). HCMV-infected cultures contained single nuclei and multinucleated GFP-positive cell bodies (Fig. 7B), indicating syncytium formation regardless of nitric oxide exposure. We attempted to quantify progenitor, neuron, and glial populations using immunofluorescence, since our goal was to evaluate changes in marker expression. However, we were unable to use this approach due to culture density and overlapping projections (Fig. 7A).
To overcome this limitation, we switched to flow cytometry to quantify differentiated neural populations, which also allowed us to identify double- and triple-marker-positive cells (68). As described above, NPCs were plated from dissociated iPSC-derived NPC neurospheres and completed using different NPC passages. The passage numbers are presented as iPSC/NPC neurosphere passage numbers. Cells from passage 45/22 (P45/22), P61/12, and P66/9 were used for these studies. NPCs were HCMV or mock infected, and flow cytometry was performed at 11 dpi. Cells were labeled with Nestin-, Tuj1-, and GFAP-conjugated fluorescent antibodies. Gates were placed to isolate live, single-cell events, and GFP+ events were identified by first gating on the unstained, uninfected population (Fig. 8A). Forward and side scatter gates were extended to include the larger GFP+ cells. It is possible that syncytia were lost despite the increased gate size; however, we reasoned that these cell bodies would be equally lost in all conditions. Uninfected cells had no GFP+ events. GFP+ events varied in infected cultures depending on the passage of NPCs, despite infection at the same MOI (Fig. 9A). For cells from P45/22, 54.7% of the live, single-cell events were GFP+. In P61/12 and P66/9 cells, GFP+ events were 1.6 and 5.7%, respectively. As GFP levels varied between experiments, we grouped GFP+ and GFP− events to analyze the whole culture population for each experiment. We used fluorescence minus one (FMO) controls (Fig. 8B) to define Tuj1/GFAP, Tuj1/Nestin, and GFAP/Nestin populations, and our gating strategy is shown in Fig. 8C.
We first determined frequencies of Tuj1+ Nestin− GFAP− events, which represent neuron populations, and Tuj1+ Nestin+ GFAP− events, which likely represent an immature neuron population, as they express both Tuj1 and Nestin (Fig. 9B and C). We quantified these populations by gating on live, single-cell, Tuj1+ events from Tuj1/GFAP populations (Fig. 8C, Q1). Tuj1+ Nestin− GFAP− (neurons) frequencies in the mock-infected vehicle-treated control groups varied between passage number. For P45/22 cultures, the frequency of Tuj1+ Nestin− GFAP− (neurons) events was 25% compared with 45% and 41% quantified in P61/12 and P66/9 cultures, respectively (Fig. 9B and C). This variation indicates intrinsic differences in the spontaneous differentiation capacity of the NPCs, and we suspect that these differences also contribute to the variability in susceptibility to infection (Fig. 9A). Upon exposure to nitric oxide, we observed a substantial decrease in the percent of Tuj1+ Nestin− GFAP− (neurons) events in P45/22 cultures in mock- and HCMV-infected populations (Fig. 9C). Less dramatic reductions were observed in P61/12 and P66/9 (Fig. 9C), suggesting that sensitivity to nitric oxide is influenced by intrinsic differences between passages and developmental states. The percent of Tuj1+ Nestin+ GFAP− (immature neurons) events for mock-infected, vehicle-treated cells were more similar between P45/22 and P66/9 at a respective 31% and 26% (Fig. 9B and C). This population was increased in P45/22 and P66/9 cultures during nitric oxide exposure regardless of infection (Fig. 9C). For P61/12, the percent of Tuj1+ Nestin+ GFAP− (immature neurons) events for mock-infected, vehicle-treated cultures was 42% and no differences were observed during nitric oxide exposure (Fig. 9C). Again, these data suggest that differentiation capacity between cell passages influences their susceptibility to nitric oxide. Cells exposed to DETA (spent donor) had similar frequencies of Tuj1+ Nestin− GFAP− cells (neurons). For P45/22 and P66/9, there was an increased percent of Tuj1+ Nestin+ GFAP− (immature neurons) compared to vehicle, though not to the level of nitric oxide-exposed cells (Fig. 9C). Overall, the results suggest that nitric oxide limits differentiation of immature neurons (Tuj1+ Nestin+ GFAP−) to neurons (Tuj1+ Nestin− GFAP−).
We next examined the percent Tuj1+ Nestin+ GFAP+ and Tuj1+ Nestin− GFAP+ cells (Fig. 9D) relative to live, single cells by gating on Tuj1+ GFAP+ events (Fig. 8C, Q2). Tuj1+ Nestin+ GFAP+ populations likely represent progenitor-like cells (69, 70). The identity of Tuj1+ Nestin− GFAP+ populations is unknown, but they may represent cells in transition to a neuron or glial state after downregulation of Nestin (69). Frequencies of Tuj1+ Nestin+ GFAP+ (progenitor-like) events varied between passages for mock-infected, vehicle-treated cultures with levels being most similar between P45/22 and P66/9 (Fig. 9B and D). We observed an increase in the P66/9 progenitor-like population in uninfected nitric oxide exposed cultures that was not observed in other passages (Fig. 9D). P61/12 had substantially less Tuj1+ Nestin+ GFAP+ (progenitor-like) events in mock-infected vehicle-treated cultures (Fig. 9B and D). DETA (spent donor) treatment decreased frequencies of Tuj1+ Nestin+ GFAP+ events in P45/22 and P66/9, suggesting some impact of nitric oxide oxidation products on this progenitor-like population (Fig. 9D). Tuj1+ Nestin− GFAP+ (transition state) populations were also variable between passages in mock-infected vehicle-treated cultures at 15.3%, 2.2%, and 5.2% for P45/22, P61/12, and P66/9, respectively (Fig. 9B and D). This population was reduced during infection of P45/22, and nitric oxide exposure also reduced these frequencies regardless of infection (Fig. 9D). Overall, our data indicate a limited impact of nitric oxide on Tuj1+ Nestin+ GFAP+ (progenitor-like) and Tuj1+ Nestin− GFAP+ (transition state) populations.
Percent of Tuj1− Nestin− GFAP+ cells, representing glial populations, were obtained by gating on Tuj1− GFAP+ (Fig. 8C, Q3) and then gating on Tuj1− Nestin− GFAP+ (Fig. 9E). We again observed differences in the frequency of glial populations between the passages of mock-infected, vehicle-treated groups with a respective 4.8%, 0.2%, and 2.3% for P45/22, P61/12, and P66/9 (Fig. 9B and E). For P45/22 and P66/9, Tuj1− Nestin− GFAP+ (glial) populations in infected, vehicle-treated cultures were modestly decreased compared with mock-infected, vehicle-treated cultures. We observed a limited impact from the spent donor on this population (Fig. 9E), suggesting that oxidation products do not alter glial populations. Nitric oxide decreased Tuj1− Nestin− GFAP+ (glial) frequencies regardless of infection (Fig. 9E), suggesting that nitric oxide limits glial differentiation. Frequencies of Tuj1− Nestin+ GFAP+ cells, which may represent immature glial cells, were below 1% and not included in further analysis.
Finally, levels of Tuj1− Nestin+ GFAP− cells, representing progenitor populations (Fig. 9B and F) were obtained by gating on Tuj1− GFAP− events and Nestin+ events (Fig. 8C, Q4). Levels of Tuj1− Nestin+ GFAP− (progenitor) cells were notably higher in P66/9 than P45/22 and P61/12 (Fig. 9B and F). Nitric oxide decreased progenitor populations in P66/9 mock-infected vehicle-treated cultures from 7.1% to 3%, with a similar reduction observed in infected cultures (Fig. 9F). The frequencies of progenitor populations for P45/22 and P61/12 were too low to observe differences (Fig. 9F). Taken together, our data suggest that nitric oxide, and to some degree oxidation products, disrupts NPC differentiation of both neuron (Tuj1+ Nestin− GFAP−) and glial (Tuj1− Nestin− GFAP+) populations with a trend toward increased immature neurons (Tuj1+ Nestin+ GFAP−) regardless of infection.
Nitric oxide is a selectively reactive free radical produced as a component of the innate immune response and has antiviral activity against HCMV replication. Prior to our studies, the impact of nitric oxide on the developing brain during HCMV infection was unknown. We modeled this complex relationship using 3-dimensional cortical organoids that recapitulate many aspects of fetal brain development. We observed that nitric oxide disrupts HCMV replication in cortical organoids. However, nitric oxide substantially disrupts tissue organization and structure in both mock- and HCMV-infected organoids (Fig. 2 and 3). Gabrielli et al. (7) observed that brains of congenitally infected fetuses have numerous necrotic brain regions with infiltrating immune cells that include macrophages and microglia that are known to produce nitric oxide (24, 25). These observations suggest that nitric oxide production during congenital infection is likely highest at the sites of infection with surrounding uninfected cells exposed to a gradient of nitric oxide. It is likely that as the nitric oxide gradient decreases, there are less pathogenic consequences for uninfected cells. In our studies, the outer layer of the organoid is exposed to the highest concentration of nitric oxide and interior cells are likely exposed to a concentration gradient. While this does not fully mimic conditions during infection, it does subject infected and uninfected to cells to various concentrations of nitric oxide, as would occur during infection. To better recapitulate physiological conditions, cortical organoids can be generated to include microglia, which produce nitric oxide via NOS2 during infection (25, 71). Studies by Ormel et al. (71) demonstrated that organoids are capable of producing mesodermal progenitors that differentiate to cells expressing classical microglial markers. The addition of microglia to the cortical organoid model of HCMV infection will allow future investigation into the role of other immune molecules, such as cytokines, on developing cortical tissue.
Microcephaly, or smaller head and/or brain size, is associated with disruption of progenitor cell populations (54). HCMV infection can result in microcephaly, which contributes to cognitive defects in congenitally infected infants (2). Dysregulation of progenitor populations during HCMV infection is thought to contribute to this birth defect (21–23). Our studies suggest that nitric oxide may also contribute to microcephaly, as exposure decreased SOX2, the regulator of progenitor maintenance, and altered neural populations, including decreased neurons (Tuj1+ Nestin− GFAP−) and glial (Tuj1− Nestin− GFAP+) cells (Fig. 5C and 9C to F). Further, cortical organoids exposed to nitric oxide had significant disruption of tissue organization and rosette structures, suggesting dysregulated progenitor populations (Fig. 2 and 3).
The severity of brain damage in congenitally infected fetal brains is associated with various factors, including viral load and inflammatory response (7). Brains with higher viral load and infiltration of inflammatory cells correlate with more severe damage than those with lower levels. During our studies, infection of NPCs resulted in variability in the frequencies of GFP+ cells per passage despite infecting at the same MOI (Fig. 9A). These differences per passage extended to the impact of nitric oxide. We speculate that intrinsic differences in NPC passage impacted susceptibility to infection and nitric oxide. Differentiation variability and low frequency of infected cells are caveats to our current study; however, we propose that low-GFP+-cell infections mimic some aspects of congenital infection, as the range in viral load impacts disease severity. Further, investigating differences in differentiation capacity is an important area for future research as these mechanisms likely have a role in disease susceptibility within the developing brain.
Metabolic reprogramming is essential for neuronal differentiation (67, 72–74). Agostini et al. (72) and Zheng et al. (67) demonstrated that basal and maximal OCRs are increased in differentiated neurons compared to progenitor cells. Zheng et al. (67) also showed that a reduction in glycolytic enzymes is essential for neuron differentiation. Our data demonstrate that nitric oxide significantly decreases maximal respiration of NPCs and reduces neuron (Tuj1+ Nestin− GFAP−) populations during NPC differentiation in both uninfected and HCMV-infected cultures (Fig. 6E and F and 9C). It is possible that nitric oxide limits the potential of NPCs to differentiate by limiting the capacity of the cell to respond to increased energy demands and forcing glycolysis to compensate for limited energy. Further, dysregulation of metabolism by HCMV infection is a plausible mechanism for the reduction in neuron populations observed during HCMV infection (13, 16). It is also possible that nitric oxide-mediated cell death results in decreased mature neuron populations. Nitric oxide can induce neuron death through several mechanisms, including inhibition of mitochondrial respiration, altered metabolism, poly-ADP-ribose polymerase (PARP) activation, and/or formation of reactive nitrogen species (75). Future studies will elucidate the mechanisms of limited neural differentiation versus cell death during HCMV infection and nitric oxide exposure.
Mitochondrial respiration generates ATP through the proton gradient established by the electron transport chain (ETC). The electron donors NADH and FADH2 are critical contributors to mitochondrial respiration as they donate electrons to complex I and II, respectively, of the ETC. HCMV infection increases mitochondrial biosynthesis, and functional mitochondria are required for efficient viral replication (76, 77). Our previous study determined that nitric oxide inhibits HCMV replication in fibroblasts and epithelial cells by a multifactorial mechanism that includes altered metabolism and decreased mitochondrial respiration during infection (48). Here, we demonstrate that nitric oxide also inhibits HCMV spread in cortical organoids and HCMV replication in NPCs (Fig. 1 and 4). Nitric oxide may have a similar mechanism of inhibition in NPCs as other cell types. In HCMV-infected fibroblasts, basal OCR is increased at 48 hpi but not 24 hpi (48, 77). In contrast, we observed an increase in basal OCR at 24 hpi in NPCs (Fig. 6C), suggesting differences in mitochondrial and metabolic manipulation in NPCs during infection. Nitric oxide reduced basal respiration in HCMV-infected NPCs to the level of uninfected cells. Direct inhibition of ETC enzymes could decrease basal respiration (28, 78); however, this mechanism is unlikely, as there is no effect on basal levels by nitric oxide in uninfected NPCs (Fig. 6C). More likely, nitric oxide alters metabolic intermediates that are increased during infection and required for the ETC, such as NADH and FADH2 from the TCA cycle. Chambers et al. (79) and others demonstrated that glutamine uptake and glutaminolysis are essential for infection by providing intermediates to the TCA cycle (80–82), and our previous studies suggest that glutamine diversion has a role in nitric oxide-mediated inhibition of HCMV in fibroblasts (48). If nitric oxide diverts glutamine from its primary role in NPCs during HCMV infection, it could account for decreased basal respiration, and ultimately, viral replication. Future studies will elucidate the metabolic impact on HCMV-infected NPCs during nitric oxide exposure.
In summary, congenital HCMV infection and its impact on the developing fetal brain remain a serious health concern. Cortical organoid models, which recapitulate some aspects of HCMV infection in the fetal brain, are important for elucidating mechanisms of disease. We combined this 3-dimensional model with more traditional 2-dimensional cell culture techniques to demonstrate that the free radical nitric oxide contributes to dysregulated and disorganized neural cell populations during HCMV infection despite its antiviral activity. Further, this study provides additional evidence that indirect effects of HCMV infection such as nitric oxide production are potent contributors to disease.
MATERIALS AND METHODS
Cell culture and virus.
Undifferentiated induced pluripotent stem cell (iPSC) colonies, initially derived from fibroblasts (Coriell fibroblast line GM003814) (52, 53), from a healthy individual were maintained in Essential 8 medium (Thermo Fisher Scientific) and grown under feeder-free conditions on Matrigel (Corning). Neural progenitor cells (NPCs) were differentiated and maintained as neurospheres (60) in Stemline (Millipore Sigma) supplemented with 0.5% N-2 supplement (Thermo Fisher Scientific), 100 ng/mL epidermal growth factor (EGF) (Miltenyi Biotech), 100 ng/mL fibroblast growth factor (FGF; Stem Cell Technologies), and 5 μg/mL heparin (Millipore Sigma). Neurospheres were dissociated using TrypLE and plated at 500,000 to 600,000 cells per well on Matrigel-coated dishes. Plated NPCs were differentiated in neurobasal medium (Thermo Fisher Scientific) supplemented with 2% B-27 (Thermo Fisher Scientific).
TB40/EFb-eGFP stocks were obtained by transfecting a bacterial artificial chromosome (BAC) encoding HCMV strain TB40/E-eGFP and a plasmid encoding UL82 into MRC-5 fibroblasts using electroporation at 260 mV for 30 ms with a 4-mm-gap cuvette and a Gene Pulser XCell electroporation system (83–86). TB40/EEpi-eGFP stocks were obtained by infecting ARPE-19 cells with TB40/EFb-eGFP. Viral medium was collected and pelleted through a sorbitol (20% sorbitol, 50 mM Tris-HCl [pH 7.2], 1 mM MgCl2) cushion at 20,000 × g for 1 h in a Sorvall WX-90 ultracentrifuge and SureSpin 630 rotor (Thermo Fisher Scientific). Viral stock titers were determined on MRC-5 (TB40/EFb-eGFP) or ARPE-19 (TB40/EEpi-eGFP) cells in 96-well dishes using a limiting dilution assay (50% tissue culture infective dose [TCID50]). GFP-positive wells were determined at 2 weeks postinfection, and resulting titers were reported in infectious units (IU) per milliliter. MOI of 0.05 or 3 IU per cell were used for infection of NPCs using TB40/EEpi-eGFP. Day 35 cortical organoids were infected using an MOI of 500 IU/μg using TB40/EFb-eGFP.
Cortical organoid cultures were differentiated from the healthy iPSC cell line according to the specification of the cerebral organoid kit from Stemcell Technologies (no. 08570) that relies on an established protocol (54). Briefly, iPSCs were plated at 9,000 cells per well onto 96-well ultralow-attachment plates for embryoid body (EB) formation and grown in EB formation medium (Stemcell Technologies). At day 5, the induction of neural epithelium was initiated by moving the EBs into an ultralow-attachment 24-well plate and feeding with induction medium (Stemcell Technologies). On day 7, neural tissues were embedded in Matrigel droplets, moved to ultralow-attachment 6-well plates, and fed expansion medium (Stemcell Technologies). Day 10 organoids were transferred to a rocker to elicit the circulation of nutrients and prevent organoids from sticking to the dish. The organoids were expanded and fed with fresh maturation medium every 3 to 4 days. This method is also described in reference 21. Organoid viability was assessed by trypan blue exclusion. Cortical organoids were incubated in 500 μL of Accutase (Thermo Fisher) at 37°C for 15 min and washed with phosphate-buffered saline (PBS) (Invitrogen) to remove residual enzyme. Organoids were dissociated by pipetting until a single-cell suspension was achieved and quantified with a Countess automated cell counter (Thermo Fisher).
Stocks of 50 mM diethylenetriamine NONOate (DETA/NO) (Cayman Chemicals) in 0.01 M NaOH were maintained at −20°C. Spent donor (DETA) was prepared by diluting DETA/NO to the appropriate concentration in medium and incubating at 37°C for approximately 72 h and then maintained at 4°C. DETA/NO cytotoxicity was assessed by plating 500,000 NPCs/well on Matrigel (Corning) coated 6-well dishes and treated with a range of concentrations or vehicle control (NaOH). Treatment was replaced every 24 h, and cell viability and number were quantified at 96 h posttreatment using a hemocytometer and trypan blue exclusion. For the remaining experiments, 200 μM and 400 μM DETA/NO were used to treat NPCs and cortical organoids, respectively.
Virus titers were determined by collecting supernatants from infected cultures at 96 hpi and plating serial dilutions on ARPE-19 cells in 12-well dishes. Cells were washed at 2 hpi and stained at 144 hpi with anti-IE1 antibody, and IE1-postive plaques were quantified. Titers of cell-free and cell-associated virus were determined by collecting supernatants and cells from the same culture. Cells from infected cultures were resuspended in 1 mL of Dulbecco’s modified Eagle medium (DMEM), 7% fetal bovine serum (FBS), and 1% penicillin-streptomycin and freeze-thaw lysed by incubating in a dry ice/ethanol slurry for 3 min followed by 10 min at 37°C. This process was repeated twice before centrifugation and collection of the supernatant. Serial dilutions of culture or lysed cell supernatants were plated on ARPE19 cells in 12-well dishes, washed at 24 hpi, and stained at 96 hpi with anti-IE1 antibody, and IE1-positive cells were quantified.
Analysis of nucleic acid and protein.
Quantitative PCR (qPCR) was used to quantify viral DNA levels. Cells were collected by trypsinization, and DNA was isolated using the DNeasy blood and tissue kit (Qiagen). Primers for HCMV UL123 (5′-GCCTTCCCTAAGACCACCAAT-3′ and 5′-ATTTTCTGGGCATAAGCCATAATC-3′) and cellular TP53 (5′-TGTTCAAGACAGAAGGGCCTGACT-3′ and 5′-AAAGCAAATGGAAGTCCTGGGTGC-3′) (Integrated DNA Technologies) were used for quantitation. qPCR was completed using SYBR green PCR mix (Thermo Fisher) and QuantStudio 6 Flex real-time PCR. Relative HCMV UL123 was normalized to cellular TP53.
Protein levels were determined by Western blot analysis. Cells were collected by trypsinization, resuspended in lysis buffer (50 mM Tris-HCl [pH 8.0], 150 mM NaCl, 1% SDS) with protease and phosphatase inhibitor, and lysed by sonication. Protein concentrations were determined with a Pierce bicinchoninic acid (BCA) assay kit (Thermo Fisher), and 20 μg of protein was resolved by SDS-PAGE using 4-to-20% gradient gels (Bio-Rad). Proteins were transferred to nitrocellulose membrane (GE Healthcare Life Sciences) using a semidry Trans-Blot transfer system (Bio-Rad), and membranes were blocked for 1 h in 5% milk in PBS-T (phosphate-buffered saline, 0.1% Tween 20) or TBS-T (Tris-buffered saline, 0.1% Tween 20). After blocking, membranes were incubated with primary antibody diluted in 5% milk or bovine serum albumin (BSA) in PBS-T or TBS-T overnight at 4°C. Membranes were washed in PBS-T or TBS-T and incubated with StarBright Blue secondary antibody (Bio-Rad) diluted in 5% milk in PBS-T for 30 min at room temperature in the dark and imaged using a Bio-Rad ChemiDoc imager.
The following antibodies and dilutions were used for Western blot analysis: mouse anti-UL44 (1:10,000; Virusys), rabbit anti-cyclin B1 (clone D5C10, 1:1,000; CST), mouse anti-p21 (clones CP36 and CP74, 1:1,000; Millipore Sigma), and rabbit anti-SOX2 (1:1,000; Millipore Sigma). The HCMV IE1 (clone 1B12, 1:1,000), IE2 (clone 3A9, 1:1,000) and pp28 (clone 10B4-29, 1:1,000) antibodies were generously provided by Tom Shenk (Princeton University). The secondary antibodies StarBright Blue goat anti-mouse and anti-rabbit (Bio-Rad) were used at a 1:10,000 dilution.
Mitochondrial stress assay.
A Seahorse XFe96 analyzer was used to determine oxygen consumption rates (OCRs). For NPCs, 50,000 cells/well were plated on Matrigel (Corning)-coated Agilent Seahorse 96-well dishes and incubated for 3 days. Cells were infected with HCMV and treated as described in the figure legends. The mitochondrial stress analysis was performed at 1 dpi, with the medium being replaced 1 h prior with XF DMEM base medium. The assay was performed by sequential injections of oligomycin (1 μM), FCCP (1.5 μM), and rotenone/antimycin A (1 μM) at the indicated times. Wells that did not respond to injection and edge wells were omitted from the analysis.
On Matrigel-coated six-well dishes, 600,000 cells/well were plated and incubated for 3 days. Cells were infected with HCMV at an MOI of 0.05 IU/cell, washed at 2 hpi, and treated with 200 μM DETA/NO, DETA (spent control), or vehicle control. Treatment was changed every 2 days. At 11 dpi, cells were collected by trypsinization, washed with PBS, and fixed and permeabilized using a cell fixation and permeabilization kit (Abcam) at room temperature Cells were incubated with conjugated antibody for 30 min at room temperature in the dark and strained through a 100- or 40-μm filter. Data were acquired on a BD LSR II flow cytometer (BD Biosciences). Cells were gated as described in the figure legends.
For flow cytometry, the following antibodies were used: mouse anti-Nestin conjugated to V450 (clone 25, 2.5 μL; BD Biosciences), mouse anti-GFAP conjugated to phycoerythrin (PE) (clone 1B4, 2.5 μL; BD Biosciences), and mouse anti-beta-tubulin class III (clone Tuj1, 15 μL; BD Biosciences).
For organoid images, cortical organoids were fixed in 4% paraformaldehyde overnight at 4°C, washed with PBS, placed in 30% sucrose in PBS for 2 to 3 days, and then transferred to optimum cutting temperature (OCT) compound. Organoids were embedded in a cryosectioning mold using OCT compound (Thermo Fisher), frozen on dry ice, and cryosectioned into 20-μm slices. Sections were immediately mounted and allowed to dry. Sections were blocked in 5% donkey serum and 0.1% Triton X-100 in PBS for 30 min, incubated in primary antibody overnight at 4°C, and incubated in secondary antibody for 1 h at room temperature. Hoechst was used to stain nuclei. Images were acquired on an upright TS100 Nikon fluorescence microscope, and NIS Elements was used for imaging and analysis.
For NPCs, 600,000 cells/well were plated on a Matrigel-coated six-well dish containing 12-mm coverslips and incubated for 3 days. Cells were infected with HCMV at an MOI of 0.05 IU/cell, washed at 2 hpi, and treated with 200 μM DETA/NO, DETA (spent control), or vehicle control. Treatment was changed every 2 days. At 11 dpi, cells were fixed in 4% paraformaldehyde for 20 min and permeabilized in 0.1% Triton X-100 for 15 min. Cells were blocked in 5% normal goat serum (Millipore-Sigma) in 0.2% Triton X-100 for 1 h and incubated in primary antibody diluted in 5% normal goat serum in 0.1% Triton X-100 overnight at 4°C. Cells were washed with PBS-T and incubated for 1 h with the appropriate secondary antibody conjugated to Alexa Fluor 568 (1:1,000) and diluted in 5% normal goat serum in 0.1% Triton X-100 at room temperature. Cells were stained with Hoechst (1:1,000) for 10 min at room temperature in the dark. Coverslips were placed on glass slides and mounted with ProLong glass antifade mountant. Images were acquired on a Nikon Ti Eclipse inverted microscope, and NIS Elements was used for imaging and analysis.
The following antibodies were used for immunofluorescence: mouse anti-Nestin (1:600; Abcam), rabbit anti-GFAP (1:1,000; Agilent), mouse anti-Tuj1 (1:100 to 1,000; Millipore-Sigma), rabbit anti-SOX2 (1:100; Millipore Sigma), anti-mouse Alexa Fluor 568 (1:1,000; Thermo Fisher), anti-rabbit Alexa Fluor 568 (1:1,000; Thermo Fisher), anti-mouse Alexa Fluor 647 (1:250), and Hoechst (1:1,000; Thermo Fisher). A TUNEL assay was performed according to the manufacturer’s instructions (Thermo Fisher).
Statistical significance was determined using GraphPad Prism application software and the appropriate statistical tests as indicated in the figure legends. P values of <0.05 were considered significant.
We thank Tom Shenk for providing antibodies against HCMV IE1, IE2, and pp28 proteins; Monika Zielonka in the Redox & Bioenergetics Shared Resource facility; Benedetta Bonacci in the Versiti-Blood Research Institute Flow Cytometry Core; and MCW Cancer Center and Children’s Research Institute’s Flow Cytometry Core. We thank Melissa Whyte and Halli Miller for their helpful advice on flow cytometry, Michele Battle for review of the manuscript, and members of the Terhune and Hudson laboratories for their input on the project.
Research reported in this publication was supported by the National Institute of Allergy and Infectious Diseases division of the National Institutes of Health under award number R01AI083281 (S.S.T.) and R01AI132414 (S.S.T. and A.D.E.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
R.L.M., A.D.E., and S.S.T., conceptualization and methodology; S.S.T., A.D.E., funding acquisition and supervision; R.L.M., B.S.O., J.W.A., S.R., investigation; R.L.M., B.S.O., J.W.A., S.R., M.L.S., A.D.E., S.S.T., formal analysis; R.L.M., M.L.S., and S.S.T., visualization; R.L.M., writing—original draft; R.L.M., A.D.E., and S.S.T., writing—review & editing.
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