New Results

Cellular events of acute, resolving or progressive COVID-19 in SARS-CoV-2 infected non-human primates

MD Fahlberg, RV Blair, LA Doyle-Meyers, CC Midkiff, G Zenere, KE Russell-Lodrigue, CJ Monjure, EH Haupt, TP Penney, G Lehmicke, BM Threeton, N Golden, PK Datta, CJ Roy, RP Bohm, NJ Maness, T Fischer, J Rappaport, M Vaccari
doi: https://doi.org/10.1101/2020.07.21.213777

Abstract

We investigated the immune events following SARS-COV-2 infection, from the acute inflammatory state up to four weeks post infection, in non-human primates (NHP) with heterogeneous pulmonary pathology. The acute phase was characterized by a rapid migration of CD16+ monocytes from the blood and concomitant increase in CD16+ macrophages in the lungs. We identified two subsets of interstitial macrophages (HLA- DR+ CD206–), a transitional CD11c+ CD16+ population that was directly associated with IL-6 levels in plasma, and one long lasting CD11b+ CD16+ population. Strikingly, monocytes were a correlate of viral replication in bronchial brushes and levels of TARC (CCL17), and worse disease outcomes were associated with high levels of cell infiltration in lungs and CD11b+ CD16+ macrophages accumulation. Importantly, this accumulation was long-lasting and detectable even in animals with mild or no signs of disease. Interestingly, animals with less signs of disease had a high IL-10:IL-6 ratio. Our results unravel cellular mechanisms of COVID-19 and validate NHP as models to test immune therapies.

The COVID-19 pandemic is a major public health crisis, causing a global medical emergency. There are currently no effective vaccines or drugs approved to completely treat or prevent COVID-19, and much remains unknown about the pathogenesis of the disease. COVID-19 presents with highly variable outcomes, and patients that develop symptoms exhibit a wide range of disease, spanning from mild (fever, cough, shortness of breath) to severe (dyspnea, pneumonia, rapidly progressing radiographic changes)1. It is currently estimated that up to 15% of COVID-19 patients progress to acute respiratory distress syndrome (ARDS)2, which is the major cause of death among fatal SARS-CoV-2 cases.

Studies elucidating molecular and immunological details of SARS-CoV-2 infection are underway, and many of the reported clinical observations point to the disease being at least partly the result of an excessive host response aimed to clear the virus but instead contributing to disease development3. This hypothesis is supported by studies on the closely related coronavirus SARS-CoV-14–6, and by clinical reports showing elevated levels of the proinflammatory cytokine IL-6 in SARS-CoV-2 infected patients, particularly among those with severe symptoms3, 7. Interestingly, the increase in IL-6 coincides with the upregulation of chemokines responsible for myelopoiesis and monocyte recruitment to the lungs3, 8 suggesting that peripheral inflammatory monocytes and tissue macrophages may play a role in the cytokine storm seen in severe COVID- 19 patients9. The underlying inflammatory state and increased myelopoiesis seen in older individuals may be responsible for the high mortality rate seen in this group10. A better understanding of immunopathology of COVID-19 will aid the development of targeted immunotherapies and effective vaccines.

Non-human primates (NHP) have been used as animal models to study acute respiratory diseases such as SARS and MERS11. In these studies, African green monkeys (AGM) have been shown to be more susceptible to severe disease than rhesus macaques (RM)12. Regardless of species differences, and given the overlap between NHP and with human immune systems, both NHP species have been useful models to study host responses to coronaviruses55. We and others have shown that both AGM and RM can be successfully infected with SARS-CoV-2 via the intranasal and the intratracheal route, or by aerosol13–16. Following exposure, AGM and RM have high levels of virus replication in the respiratory tract for up to two weeks, and both species develop pulmonary pathology characteristic of COVID-19, that varies from very mild to severe17. Other groups have described the very early immune events in SARS-CoV-2 infected rhesus macaques, with perturbation of monocytes populations, increased macrophages in lungs, and cellular activation in blood all happening a few days following the infection18.

Here, we extend on these findings by describing the kinetics of immune events following infection, spanning from the acute phase up to four weeks following infection, in the blood and the lungs of four African green monkeys (AGM) and four rhesus macaques (RM) with a heterogeneous spectrum of COVID-19. Overall, we observed a multiphasic mobilization of innate and adaptive cells from the blood to the lung that is strikingly common to both species. The acute inflammatory phase was characterized by an increase in the frequency of monocytes that was positively associated with bronchial viral replication. Consistent with the resolution phase of inflammation in the lung19, the second phase was characterized by a switch toward type 2 responses in both species. We identified two populations of interstitial macrophages (HLA- DR+ CD206–) in BAL; 1) CD11c+ CD16+ population increasing during the acute phase of infection that was associated with IL-6 levels in the plasma, and 2) a long lasting CD11b+ and CD16+ population accumulating in lungs of both AGM and RM, particularly in those with increased disease severity. Animals with more pronounced clinical signs had a higher level of infiltrates in lungs when compared with animals with no or mild signs, and higher ratio of IL- 6:IL-10 (pro-inflammatory: anti-inflammatory) cytokines in plasma.

By investigating temporal and spatial changes in immune cells our study links together findings in both NHP and human patients and unravels new cellular events of COVID-1920, 21. Further, we validate the use of NHP as a model to study immunopathology and to test immune therapies and vaccines for SARS-CoV-2.

Results

Increased levels of monocytes and chemokines after infection is associated with viral load and disease severity

Four adult RM and four adult AGM were exposed to SARS-CoV-2 by either aerosol or mucosal challenge including buccal, intranasal, intratracheal, and conjunctival (multi-route) exposure (Supplementary Table 1). The study concluded at 4 weeks post exposure (Fig. 1A)16. All eight animals had detectable viral load by RT-PCR on nasal and pharyngeal swabs and bronchial brushes by week 1 (Fig. 1B, Supplementary Table 2). Viral RNA was also detected in pharyngeal, nasal, buccal, rectal and vaginal swabs. No differences in viral load (VL) were observed between the different routes of exposure or between the two species at week 1. Of note, by week 2 AGM had higher VL than RM in the bronchial brushes and by week 3 VL in RM was mostly undetectable while it remained above detection levels in AGM until study completion (Fig. 1B and Supplementary Table 2). Similar to humans, the course of disease varied widely across animals, as detailed in Table 1. Of the four AGM enrolled, two animals developed severe respiratory signs and were euthanized on days 8 (NC34) and 22 (NC33) post-infection, following established endpoint criteria for NHP study protocols, and as we previously reported16. Of the two remaining AGM, NC38 had multifocal mild to moderate interstitial pneumonia scattered throughout all lung lobes and NC40 was mostly asymptomatic and had scant inflammation in all lung lobes at necropsy. Three of the four RM (GH99, HD09, and FR04) developed pneumonia characterized by a granulomatous to pyogranulomatous inflammatory response. This was severe in GH99, mild in HD09, and minimal in FR04. Histopathology of the fourth RM (HB37) revealed a lymphocytic vasculitis and proliferative vasculopathy in the right middle lung lobe. Viral load was not significantly associated with pulmonary pathology, however we observed an inverse association between viral load at week 1 and disease severity ranking (1 to 8 from most severe to least severe) based on the above observations and histopathology findings (AGM: R = 0.8; RM: R = –0.4, all animals: P = 0.057, R = – 0.71, by the Spearman correlation test) (Fig. 1C and SF 1A and 1B). Clinical signs observed during the course of the study included mild tachypnea (day 11) and cough (day 15) in GH99. Intermittent mild to moderate pyrexia was observed in HB37 (day 1-21 post infection). Radiographic changes included a nodule in the right caudal lung lobe (day 11) in GH99, and HD09 also developed a mild, focal increased opacity in the ventral lung field at day 11 (data not shown).

Figure 1.
Figure 1. Monocytes are a correlate of bronchoalveolar viral replication.

(A) Schematic of study design. Arrows represent time of inoculation with SARS-CoV-2 (day 0) or sacrifice (up to 28 days post infection). Unless stated otherwise, rhesus macaque data are shown in teal, and African green monkey data are shown in magenta. (B) SARS-CoV-2 RNA levels in bronchial brushes over time. (C) Association between SARS-CoV-2 load in bronchial brushes and ranking of disease severity in all eight animals (most severe case = 1; mildest case = 8). (D) Changes in monocyte count in blood before and after infection. (E) Proportion of total monocytes (HLA- DR+ CD16+ and/or CD14+ out of CD45+ leukocytes) in PBMCs over time. (F) Correlation between SARS-CoV-2 viral load in bronchial brushes and absolute count of monocytes in blood at one-week post-infection (1wk pi). (G) Representative flow cytometry changes in non- classical monocytes after the infection in blood (1wk pi). (H) Percent of classical (CD14+ CD16–) and (I) non classical monocytes (CD14– CD16+) out of total monocytes in PBMCs. (J) Correlation between monocytes absolute numbers in blood and TARC (CCL17) in plasma (1wk pi). (K) Relationship between fold change of chemokines and SARS-CoV-2 viral load in bronchoalveolar brushes (1wk pi). (L) Heatmap displaying log2 fold-change of 8 chemokines involved with traffic of monocytes to the lung. (1wk pi). Animals are ordered by ranking of disease severity.

View this table:
Table 1: Heterogeneous clinical and pathological outcomes following SARS-CoV-2 infection

Interestingly, virus-driven immunological changes were remarkably similar between species at this time (Fig. 1 and SF 1). To investigate overall changes driven by viral replication in both groups, we combined their immunological analysis, however at the same time we maintained different specifications for each species (teal = RM and magenta = AGM). Data and statistics are also shown separately per group in Supplementary Figure 1. At 1 week after infection both AGM and RM had an increase in the absolute number and frequency of monocytes in the blood (P = 0.065, and P = 0.02 respectively, by the Tukey multiple comparison test, n = 8) compared to baseline levels (Fig. 1D and 1E). This change was significant in AGM alone (absolute number: P = 0.0006, n = 4), and was also observed in all RM, particularly GH99 (SF 1C-1F). The absolute number of neutrophils did not change over time (SF 1G). Interestingly, the absolute number of monocytes at week 1 was strongly positively associated with the viral load levels in the bronchial brush samples (P = 0.002, R = 0.93 by the Spearman test) when all the animals were combined (Fig. 1F) (AGM: R = 0.8; RM R = 1, not significant in separated group, SF 1H and 1I). We then used the CD14 and CD16 markers to discriminate between three circulating monocytes subtypes: classical (CD14+ CD16–), intermediate (CD14+ CD16+) and non-classical (CD14– CD16+) within the total (HLA-DR+ CD16+ and/or CD14+) monocyte population by flow cytometry (Fig. 1G - 1I and SF 1J- 1N). At week 1, the frequency of classical monocytes was significantly increased in blood, as seen in humans22, and the non-classical population was profoundly depleted (increased levels baseline versus week 1: classical P = 0.007 and decreased non- classical P = 0.005 in all animals combined, n = 4) (Fig. 1H and 1I, and SF 1J- 1K). Pre-infection levels were restored at 3 weeks post-infection (week 1 vs. week 3: classical P = 0.03 and non-classical P = 0.03). These results suggest a rapid and robust recruitment of patrolling monocytes with antiviral activity in tissues following infection in both species. We analyzed a group of eight chemokines known to be involved in the mobilization of innate cells, including monocytes, from the blood to the lung: IP-10 (CXCL-10), MCP-1 (CCL2), MCP-4 (CCL13), Eotaxin, TARC (CCL17), MDC (CCL22), MIP-1a and MIP-1b (SF 2A-2H). Some of the cytokines measured at baseline in plasma differed between the two species (MCP-4, Eotaxin, MIP-1b and MDC, Supplementary Table 3 and SF 2I). Plasma levels of TARC (CCL17), a ligand for CCR4 that is involved in Th2 cell recruitment23, were strongly associated with the frequency of monocytes in blood (P = 0.003, R = 0.95, Spearman test, adjusted for multiple comparison: P = 0.02) (Fig. 1J). Similarly, TARC was also associated with viral replication in BAL at week 1 (Fig. 1K and SF 2J) (P = 0.02, R = 0.857, adjusted for multiple comparisons: P = 0.2), as was IP-10 (P = 0.05, R = 0.78, adjusted for multiple comparisons) when all animals were grouped together (Fig. 1K and SF 2K), a ligand for CXCR3 that is associated with disease severity and predicts progression of COVID-19 in humans24. To gain a better understanding of a common milieu possibly driving monocyte migration observed in both groups we analyzed changes in the levels of these markers with respect to the baseline. Overall this analysis revealed that animals with the most severe disease outcomes had higher levels of chemokines at week 1, including IP-10, TARC (CCL17) (Fig. 1L).

Myelomonocytic cells infiltrate BAL and are associated with IL-6

Influx of macrophages has been observed in the lung of severe cases of COVID-19 in humans25. To monitor cell trafficking into the lungs, we collected bronchoalveolar lavage (BAL) at baseline and after each week following infection, up to week 3. As expected, the majority of the cells found in BAL at baseline were alveolar macrophages (AM)26. These cells are readily identified by the mannose receptor C type 1 (CD206) marker, express high levels of HLA-DR, and are positive for the scavenger receptor CD163 (Fig. 2, SF 3A and 3B), we observed a decrease in the proportion of AM at week 1 post infection in animals of both species, corresponding with an influx of myeloid HLA-DR+ CD206– CD163– cells (Fig. 2A and 2B, SF 3C). Levels of AM similar to pre- infection were restored at week 2 in all but one animal (GH99), and a concomitant increase of CD86 mean fluorescent intensity (MFI) in this population was observed suggesting an M1 activation state27 (SF 3D). An increase was also seen in the frequency of CD206+ CD163– non- macrophages that have been previously described as endothelial cells which facilitate lymphocyte trafficking and they are detectable in BAL during inflammatory events (Fig. 2C)28.

Figure 2.
Figure 2. Myelomonocytic cells increase in BAL is associated with IL-6.

(A) Decrease of alveolar macrophages (HLA-DR+ CD206+ CD163+ cells) in the BAL after infection. (B) Increase in the frequency of myelomonocytic infiltrates (HLA-DR+ CD206– CD163–) and (C) of endothelial cells (HLA-DR+ CD206– CD163+) at infection in BAL (1wk pi). (D) tSNE plots displaying kinetics of expression of CD16, CD11b and CD11c markers on live/ HLA-DR+ cells populations in BAL before and after infection. (E) Gating of CD11c+ and CD11b+ populations on the tSNE map based on antigen expression and discrete density clustering. (F) Characterization of populations shown in (E). The heatmap depicts fold-change values of each antigen compared to channel values of negative-expressing cell populations. (G) Increase in the percentage of CD16+ HLA-DR+ cells and (H) in the percentage of interstitial macrophages (CD16+ CD206– HLA-DR+) expressing of CD11c and (I) CD11b in BAL overtime.

T-distributed stochastic neighbor embedding (tSNE) analysis revealed changes in the phenotype of CD16, CD11b, and CD11c positive populations most prominently at 1-2 weeks post infection (Fig. 2D - 2F). In contrast to what we observed in the blood, there was an increase of CD16+ HLA-DR+ cells in BAL over time (Fig. 2G, and SF 3E and 3F). We observed a transient increase in the level of CD11c+ CD16+ macrophages, which function has been described as patrolling, and they participate in rapid tissue invasion and monocyte survival (baseline vs. week 1: P = 0.015) (Fig. 2H)29, 30. The frequency of CD11c+ macrophages in BAL at week 3 was strongly associated with the levels of the proinflammatory IL-6 cytokine in the plasma at the same time point (week 3: P = 0.003, R = 0.96, Spearman test) (SF 4). The CD11b marker is required for monocyte migration to sites of inflammation31, and is highly expressed on myeloid cells recruited from the blood and differentiating into macrophages32. We also observed an increase in the percentage of CD11b+ CD16+ HLA-DR+ CD206– macrophages in BAL at week 2 and 3 following infection (Fig. 2I), a population that is involved in regulating lung inflammatory response19, 27, 33, 34. The gating strategy for the two macrophages populations is shown in Supplementary Figure 3G.

Interstitial macrophages accumulate in lungs of infected animals

Lungs were collected for all animals at the end of the study, corresponding to 4 weeks post infection (1 week following the last BAL collection shown in Figure 2), with the exception of NC33 and NC34. These two animals had higher levels of IL-6 at necropsy compared to the other six monkeys, and NC34 had symptoms similar to ARDS16 (Fig. 3). Overall, infiltration of macrophages was observed in the lungs of SARS-CoV-2 infected AGM and RM at necropsy and was more robust in animals with severe disease (Fig. 3A). We further characterized macrophages population in the lung by staining with the AM marker CD206 (in blue) and with CD11b integrin (in green) and CD16 (red) (Fig. 3B). Low numbers of macrophages were seen in the lungs of a non-infected control RM, scattered throughout the interstitium and alveolar spaces, as shown by the arrows. In the control this population was bi-morphic and composed of CD11b negative CD16+ CD206–(arrow) in the alveolar septa or CD16+ CD206+ cells within the alveolar spaces (arrowhead) (Fig. 3B). In all animals that were infected we could detect CD11b+ CD16+ macrophages. FR04 (RM ranking = 7) lungs had macrophages scattered throughout the interstitium and alveolar spaces. The phenotypic population was heterogeneous and contained low numbers of CD11b+ CD16+ CD206– macrophages. The most severe RM, GH99 (ranking = 3), had large numbers of macrophages multifocally surrounding small airways denoted by the asterisks. Of note this animal had the higher absolute number of monocytes in the blood at week 1 (SF 1D). Low numbers of CD11b+ CD16+ CD206– cells were scattered throughout the predominant population of CD11b+ CD16+ CD206+ macrophages. Macrophages were also scattered throughout the interstitium and alveolar spaces. NC34 and NC33 (AGM ranking = 1 and 2 respectively) had multifocal to coalescing areas of hemorrhage (NC33 indicated by the arrows), and pulmonary parenchyma infiltrated by multifocal aggregates of macrophages (NC34) with infiltration by low to moderate numbers of macrophages. The phenotypic population was heterogenous and composed of moderate numbers of CD11b+ CD16+ CD206– macrophages. Altogether these findings present a picture of long-lasting lung inflammation that, albeit more pronounced in animals with severe signs of disease, it is also present in animals with no detectable viral load and mild or no signs of disease after 4 weeks from the infection.

Figure 3.
Figure 3. Interstitial macrophages are increased in the lung of SARS-CoV-2 infected animals.

(A) Macrophage infiltration varied from minimal to severe in both rhesus and AGM infected with SARS-CoV-2. (B) Fluorescent immunohistochemistry in the lung of AGM and RM showing infiltration of CD16+ CD11b+ CD206– macrophages (arrows), with lesser numbers of CD16+ CD206+ macrophages (arrowheads). Inset showing a higher magnification of CD16+ CD11b+ CD206–macrophages (arrows). Asterix shows macrophages multifocally surrounding small airways in GH99. White: DAPI (nuclei); Green: CD11b; Red: CD16; Blue: CD206.

NKG2a+ cells and PD-1+ T cells decrease in blood and increase in BAL overtime

Infiltration of lymphocytes was observed in the lungs of SARS-CoV-2 infected AGM and RM at necropsy and was more robust in animals with severe disease (Fig. 4A). We first analyzed the kinetics of innate NK cells subsets, that we defined as HLA-DR– CD3– cells expressing the NKG2a receptors, which are known to traffic to the lung35 (Fig. 4B-D, and SF 5A and 5B) Interestingly, the frequencies of NKG2a+ cells significantly decreased in peripheral blood of both AGM and RM over the course of infection (baseline vs. week 3, P = 0.03) (Fig. 4B). Consistent with the pattern described for the myeloid population, we also observed a concomitant increase in the frequencies of NKG2a+ cells in BAL (baseline vs. week 3, P = 0.05) (Fig. 4C and 4D, SF 5A and 5B). Four out of four AGM had decreased NKG2a+ cells in peripheral blood, coinciding with an increase of the same cells in BAL. While the pattern of NKG2a+ cells in peripheral blood of RM was not statistically significant, nonetheless in this species there was a marked increase of those cells in the BAL. These results suggest that the loss of NKG2a+ cells in the PBMCs and the gain of NKG2a+ cells in the BAL may coincide with an egress of these cells from peripheral blood to tissues. Accordingly, the frequency of NKG2a+ cells expressing CXCR3, a marker associated with NK cell migration to the lung36 also significantly decreased in blood over time (baseline vs. week 3, P = 0.03, Fig. 4E). Despite this change in lymphocyte phenotype, the absolute number of lymphocytes did not change over the first 2 weeks of infection (Fig. 4F) though it is important to note that all 3 remaining AGM had an increase at week 3 (SF 5C).

Figure 4.
Figure 4. Lymphocytes infiltration in the lungs is associated with disease severity.

(A) Histopathologic findings showing infiltration of lymphocytes in the lung of AGM (NC40 and NC34) and RM (FR04 and GH99) with mild (NC40 and FR04), moderate (GH99), and severe disease (NC34). (B) Frequency of NKG2a+ cells in the blood or (C) BAL of 4 AGM and 4 RM at baseline, 2 weeks and 3 weeks post SARS-CoV-2 infection by flow cytometry. (D) Representative flow cytometry plot showing decreased frequency of NGK2a+ cells in PBMCs and increased frequency in BAL to baseline in at week 3 post SARS-CoV-2 infection respect to baseline. One animal for each species is shown. (E) Decrease in the frequency of CXCR3+ NKG2a+ cells in the blood at 2 weeks, and 3 weeks post SARS-CoV-2 infection compared to the baseline. (F) Absolute number of lymphocytes before and after infection. (G) tSNE plots displaying kinetics of PD-1 expression (in red) on lymphocytic cell populations over time. (H) Frequency of PD-1+ Th2 (CXCR3– CCR6–) CD4+ T cells is increased in RM and (I) in AGM overtime following the infection. (J) Changes in the percentage of Th2 (purple), Th1 (blue) and Th17 (grey) cells within the total PD-1+ CD4+ T cell population. (K) Significant increase in the percentage of PD-1+ CD8+ T cells in BAL.

tSNE analysis revealed an increase in the level of PD-1 at weeks 2 and 3 post-infection, overlapping with spatial regions corresponding to both CD4+ T and CD8+ T cells (Fig. 4G). In fact, the frequency of PD-1+ CD4+ T cells was elevated in the blood of both RM and AGM (Fig. 4H, and 4I, respectively). During the first week following the infection we observed an increase in IP-10 and IFN-γ (SF 5E), however we also observed a switch toward Th2 type responses characterized by the increase of Th2 cytokines IL-5 and IL-13 over time (SF 3E). Accordingly, a gradual increase of Th2 type cells (defined as CXCR3– CCR6 – CD4+ T cells)37 was seen in the blood within the PD-1 positive population (Fig. 4J). The increase of PD-1+ Th2 cells was significantly and positively associated with IL-4 cytokine levels (P = 0.016, R = 0.94, Spearman test) (SF 5F). Moreover, we observed an increased level of PD-1+ CD8+ T cells in blood and BAL (BAL: baseline vs. week 2, P = 0.015; baseline vs. week 3, P = 0.03) (data not shown and Fig. 4K). Animals divided by species are shown in SF 5G and 5H.

The ratio between the anti-inflammatory cytokine IL-10 and the pro inflammatory cytokine IL-6 is associated with disease progression

The pro-inflammatory cytokine IL-6 has been described as a strong predictor to disease progression in humans3, 7. We measured IL-6 and IL-10 in the plasma of all the animals (Fig. 5A). While IL-6 levels alone did not associate with disease severity in NHPs, the ratio of IL-10 to IL-6 levels measured in plasma associated with both the ranking of disease severity and with the pathological score that was obtained by analyzing the extension of the inflammation and edema of the lungs (R = 0.76 P = 0.037; and R = –0.76, P = 0.03, by the Spearman correlation test, all animals) (Fig. 5B and C). These findings show a complex series of immune events in the lung of that are most likely initiated as a host response to the virus and then evolving overtime toward less antiviral responses, as summarized in Figure 5D.

Figure 5.
Figure 5. Increased IL-6 in the absence of IL-10 is associated with disease progression.

Levels of IL-6 and IL-10 in plasma overtime (pg/ml) (A) and (B) correlation between the ratio of IL-10 to IL-6 levels measured in plasma and the ranking of disease severity. (C) Levels of IL-6 and IL-10 after infection in plasma (pg/ml) in individual animals. (D) Summary of immune events in lungs and blood of SARS-CoV-2 infected animals, from the 1-acute phase 2- switch to Th2 type responses and 3- resolution phase.

DISCUSSION

There is an urgent need to develop safe and effective strategies to prevent and treat COVID-19. The development of such strategies requires a clear understanding of the immune cells involved in COVID-19 and their unique role in pathogenesis. Clinical observations in patients and few studies on non-human primates have started to shed light on the immune aspects involved in COVID-199, 20, 38, 39. Here, we further characterized immune events following SARS-CoV-2 infection in two NHPs models recapitulating crucial aspects of COVID-19 lung disease14, 16 (Table 1). Altogether our findings provide the first evidence in both AGM and RM of a multiphasic immune response characterized by 1) an initial inflammatory phase with monocyte recruitment to the lung 2) a gradual switch from a type 1 to type 2 response, and 3) a “make it or break it” phase with either an increase in anti-inflammatory cytokines IL-10, or IL-6 a pro- inflammatory cytokines associated with disease progression in humans, as summarized in Figure 5D.

By examining the temporal and spatial distribution of monocytes and macrophage in blood and BAL and lungs, we confirmed and extended on studies in humans and NHPs suggesting increased myelopoiesis with recruitment of myeloid cells from the blood to lungs during the infection that is initially driven by the virus replicating in the lung3, 14, 20. Similar to what has been described by Wilk et al. and Thevarajan et al. in humans, we observed an increase of CD14+ monocytes, and a corresponding drop in the frequency of CD16+ monocytes in the peripheral blood20, 21. Importantly, we extend on these findings by showing a simultaneous increase in the frequency patrolling CD16+ HLA-DR+ monocytes in BAL occurring in both species and by describing the kinetics of two CD16+ HLA-DR+ CD206– macrophages populations that preferentially populate the alveolar interstitium and exhibit features of antigen-presenting cells40: CD11c+ CD16+ cells arising from intravascular CD16+ patrolling monocytes in the acute phase and associating with IL-6 levels, and a CD11b+ macrophage population known to be involved in resolution of inflammation in the lung19, 27, 33, 34. The origin and the exact function of these interstitial macrophages remains to be determined.

Differently than was observed in humans and previous reported data in RM, we did not detect an increase in the number of neutrophils as these responses may have come up earlier than 1 week after infection as shown by others18.

Common to both species was the observation that animals with higher levels of myeloid and lymphoid cell infiltration in the lung had worse disease outcomes. Several studies in both SARS- CoV-1 and CoV-2 have reported depleted peripheral NK cell counts in severe patients9, 20, 22, 38. Our data support the notion of NKG2a+ cell trafficking to the lung during SARS-CoV-2 infection, most likely mediated by IP-10 (CXCL-10), which promotes the recruitment of CXCR3+ Th1 thymocytes and NKs41, 42. Expression of NKG2a has been linked to both activation and inhibition of inflammation43; the function in SARS-CoV-2 remains to be investigated. In the periphery, we also observed increased levels of PD-1 on T cells. This is consistent with findings in humans describing of an exhaustive T cell phenotype during SARS-Cov-1 and Cov-2 infection9, 44.

For the first time, we identified TARC (CCL17) as central to myeloid recruitment in the lung during SARS-CoV-2. Both dendritic cells and CD11b macrophages in the lung have been identified as a source of CCL17 in the airways in other pulmonary inflammatory diseases45, 46. These cells promote the recruitment of T helper type 2 (Th2) lymphocytes into the airway45. Remarkably, we observed a gradual increase of the recruitment of Th2 lymphocytes into the airway Th2 cell frequency and of Th2 type cytokines such as IL-5, IL-13. These findings show a complex series of immune events in the lung of that are most likely initiated as a host response to the virus, and that they may evolve into other less effective antiviral responses as either an attempt to tissue repair/resolution of inflammation, as summarized in Figure 5D or as the result of virus direct immunomodulatory effects on host responses6.

Intrinsic species-specific susceptibility to disease progression have been described for NHP in SARS-CoV-16 with AGM being more susceptible than rhesus to severe disease. There were unexpected similarities in the kinetics and the quality of the immune responses analyzed in RM and AGM, particularly during the acute phase of infection, however AGM did not clear the infection as well as RM and two AGM had most severe symptoms47. While the number of animals enrolled in this study was not sufficient to look at species-specific differences and their effect on disease progression, we found that a higher ratio of IL-6:IL-10 cytokine in plasma was associated with disease severity. Undoubtedly, more studies are needed to prove or disprove the single contribution of IL-10 and IL-6 in the outcome of COVID-19 disease and to identify the source of these two potential key players48.

Overall, this study uncovers critical steps in the immune events of the host response to SARS- CoV-2 and describes new correlates of viral replication and disease progression. Importantly, we show viral driven inflammation/cell recruitment to the lung occurs rapidly, and lasts up to four weeks post infection, which may explain in part the observed slow recovery from COVID-19 seen humans. Finally, these findings have the potential to aid the interpretation of the results of the human trials that are currently underway for several vaccine candidates and may be instrumental in generating new targeted therapeutic strategies aiming at resolving the severe immune deregulation of the lung of COVID-19 patients.

Methods

Ethical statement on animal use and SARS-CoV-2 handling

Animals and infection. The Institutional Animal Care and Use Committee of Tulane University reviewed and approved all the procedures for this study. The Tulane National Primate Research Center is fully accredited by the AAALAC. All animals were cared for in accordance with the ILAR Guide for the Care and Use of Laboratory Animals 8th Edition. The Tulane University Institutional Biosafety Committee approved the procedures for sample handling, inactivation, and transfer from BSL3 containment.

Animals and infection

Four adult-aged African green monkeys (Caribbean origin) and four rhesus macaques (Indian ancestry) were exposed to SARS-CoV-2; 2019-nCoV/USA-WA1/2020 409 (MN985325.1). The virus stock was prepared in Vero E6 cells and the sequence confirmed by PCR and/or Sanger sequencing. Plaque assays were performed in Vero E6 cells. The rhesus macaques were from the Tulane National Primate Research Center breeding colony pathogen free for simian type D retrovirus (SRV), macacine herpesvirus 1 (B virus), simian immunodeficiency virus (SIV), simian T cell lymphotropic/leukemia virus (STLV), measles virus (MV) and tuberculosis (TB). The African green monkeys were wild caught and at the TNPRC for over a year before being assigned to this study. To mimic different possible routes of infection in humans, animals were exposed to the virus either by aerosol (inhaled dose of 2.0 x 103 and 2.5 x 103 TCID50) or by inoculating a cumulative dose of 3.61 x 106 PFU through multiple routes (oral, nasal, intratracheal, conjunctival) (Supplementary Table 1).

The multi route exposure was given to four animals, one adult RM male (GH99, 14 years old), one adult RM female (HD09, 13 years old) and two AGM, one aged male (NC40, 16 years old, approximately) and one aged female (NC33, 16 years old, approximately). An additional two adult males RM (FR04 and HB37, 15 and 13 years old, respectively) and one aged male and female AGM: NC34 and NC38 (16 years old, approximately) were exposed by aerosol (Supplementary Table 1).

Cell count and CO2 and SpO2 pressure measurements

Total white blood cell counts, white blood cell differentials, red blood cell counts, platelet counts, hematocrit values, total hemoglobin concentrations, mean cell volumes, mean corpuscular volumes, and mean corpuscular hemoglobin concentrations were analyzed from blood collected in tubes containing EDTA using a SYSMEX XN-1000v Hematology Analyzer in the Level 2 Lab and a Sysmex XS- 1000 in the Level 3 Lab. CO2 measurements were taken using the Piccolo Express Chemistry Analyzer in the Level 3 lab and is part of the Metabolic Comprehensive Panel. SpO2 values were obtained using the Masimo Rad-57 Handheld Pulse Oximeter.

Sample collection and processing

Blood EDTA was collected by venipuncture and bronchial brush and BAL by bronchoscopy. For the latter, the bronchoscope is introduced into the trachea and directed into one of the bronchi. The bronchoscope is advanced to the point where the diameter of the bronchoscope approximates the dimensions of the bronchus. The bronchial brush is passed through the channel of the bronchoscope. The brush is gently advanced until the lumen of the bronchus or bronchioles approximates the dimensions of the brush. The brush is then be gently advanced back and forth against the mucosa to gather cells. The brush is removed from the channel of the bronchoscope and the brush is placed into 200 μL of DNA/RNA Shield X1 (Cat.# R1200, Zymo Research, Irvine, CA). The bronchoscope remains in place while two aliquots of 20 ml of saline each are instilled into the bronchus and subsequently aspirated. This procedure is then repeated in the opposite lung.

Blood was layered over Ficoll-Plaque Plus (GE Healthcare17144003) and centrifuged for 30 minutes in cap-locked adaptors. Differential migration of cells during centrifugation results in the formation of layers containing different cell types and allows collection of peripheral blood mononuclear cells (PBMCs) from other populations. PBMCs were collected, counted and aliquoted in tubes or resuspended in freezing media containing Fetal Bovine Serum and DMSO. For bronchoalveolar lavage (BAL), cells and supernatant were separated by centrifugation (15 minutes at 2,000 x g at RT) in cap-locked adaptors and cells were lysed in Ammonium Chloride Potassium (ACK) lysing buffer at room temperature for 5 minutes (Gibco# A1049201) washed in 2% FBS, and resuspended in PBS and counted under laminar flux hood in BCL3. Fresh cells were stained for flow cytometry.

BAL and PBMCs staining

Freshly obtained PBMCs and cells obtained from BAL were stained with appropriate antibody cocktails and incubated for 30 minutes at room temperature. Antibodies are listed in Supplementary Table 4. Cells were fixed with 4% or 2% PFA-BSA overnight. Samples were run on a FACSAria Fusion flow cytometer (Becton Dickinson) within a Biosafety Cabinet, and each sample was run for at least five minutes. For all panels, the LIVE/DEAD Fixable Aqua Dead cell Stain kit was used (ThermoFisher #L34957). Alveolar macrophages were gated based on the size and the expression of HLA-DR, CD206 and CD163 markers. Myeloid infiltrates were gated as cells negative for CD206 and CD163 and positive for HLA-DR, CD16 and CD11b. The gating strategy is shown in Supplementary Figure 3G. NKG2a+ cells were gated on the CD3 negative parent population. Th1, Th2, and Th17 cells were gated as previously described37. tSNE analyses were performed in FlowJo using the opt-SNE algorithm and designed in R using the ggplot2 package49.

Histopathology and scoring

Tissue samples were fixed in Z-fix (Anatech), embedded in paraffin and 5 μm thick sections were cut, adhered to charged glass slides and stained with hematoxylin and eosin. All slides were scanned on a Zeiss Axio Scan.Z1 digital slide scanner. Images were acquired using HALO software (Indica Labs). Quantification of pathologic lesions was performed with HALO software. Machine learning (Random Forest) algorithms were trained by a board -certified pathologist to recognize fluidic and cellular inflammation in the lungs of the RM and AGM. Scores were assigned to fluidic and cellular inflammation, separately, based on the percentage of lung affected. Fluidic and cellular inflammation scores were summated for all lobes and animals were ranked from 1 (highest sum) to 8 (lowest sum).

Plasma cytokines and chemokines

Plasma was collected by spinning and was thawed before use. Cytokines were measured using 497 Mesoscale Discovery. IFN-γ, IL-4, IL-6, IL-8, IL-10 and IL-13 were part of the V-Plex Proinflammatory Panel 1, 10-Plex. V-plex. The chemokine panel (#K15049D, Mesoscale Discovery, Rockville, Maryland) was also used, following the instructions of the kit. The plate was read on a MESO Quickplex 500 SQ120 machine. Heatmaps were generated using the ‘pheatmap’ package in R50, 51. Data were normalized by dividing raw values at week 1 by baseline values for each animal, followed by the application of log2. Values below the limit of detection were replaced with the lowest limit of detection value based on the standard curve for each run, or with the lowest value detected during the run, whichever was smaller. Bubble plots were generated using the ‘ggplot2’ package in R, using the same normalized data shown in the heatmap49. Scatter plots were drawn using raw data points and display Pearson’s correlation coefficients and a 95% confidence interval.

RNA isolation

Swab and bronchial brush samples were collected in 200 μL of DNA/RNA Shield X1 (Cat.# R1200, Zymo Research, Irvine, CA) and extracted for Viral RNA (vRNA) using the Quick-RNAViral kit (Cat.# R1034/5, Zymo Research). The Viral RNA Buffer dispensed directly to the swab in the DNA/RNA Shield. The swab was directly inserted into the spin column. The vRNA was eluted (45μL) and 5 μL was added in a 0.1 mL fast 96-well optical microtiter plate (Cat.# 4346906, ThermoFisher, CA).

Viral RNA detection

RT-qPCR reaction TaqPath1-Step MultiplexMaster Mix was used (Cat.# A28527, ThermoFisher) along with the 2019-nCoV RUO Kit (Cat.# 10006713, IDTDNA, Coralville, IA) targeting the N1 amplicon of N gene of SARS2-nCoV19 (accession MN908947). The master mix was added to the microtiter plates covered with optical film (cat. #4311971; ThermoFisher) and then was vortexed and pulse centrifuged. The RT-qPCR program consisted of incubation at 25°C for 2 minutes, RT incubation at 50°C for 15 minutes, and an enzyme activation at 95°C for 2 minutes followed by 40 cycles of denaturing step at 95°C for 3 seconds and annealing at 60°C for 30 seconds. Fluorescence signals were detected with an Applied Biosystems QuantStudio6 Sequence Detector. Data were captured and analyzed with Sequence Detector Software v1.3 (Applied Biosystems, Foster City, CA). Viral copy numbers were calculated by plotting Cq Values obtained from unknown (i.e. test) samples against a standard curve representing known viral copy numbers. The limit of detection of the assay was 10 copies per reaction volume. A 2019-nCoVpositive control (Cat.#10006625, IDTDNA) were analyzed in parallel with every set of test samples. A non-template control (NTC) was also included.

Immunohistochemistry

5 μm sections of Formalin-fixed, paraffin-embedded lung were mounted on charged glass slides, baked overnight at 56oC and passed through Xylene, graded ethanol, and double distilled water to remove paraffin and rehydrate tissue sections. A microwave was used for heat induced epitope retrieval. Slides were heated in a high pH solution (Vector Labs H-3301), rinsed in hot water and transferred to a heated low pH solution (Vector Labs H-3300) where they were allowed to cool to room temperature. Once cool, slides were incubated with Background Punisher (Biocare Medical BP974H) for 10 minutes, washed and incubated with serum free protein block (Dako X0909) for 20 minutes. Immediately following blocking, a mouse anti-CD16 primary antibody was diluted in the serum free protein block and added to the slides for 60 minutes. Slides were washed and a MACH3 AP kit (Biocare Medical M3M532) was used to label the primary antibody. Both the MACH3 mouse probe and polymer were incubated for 20 minutes with washes in between. Signal was developed with Permanent Red substrate (Dako K0640) for 13 minutes to allow for visualization of the CD16+ cells. Slides were thoroughly washed and incubated with a blocking buffer comprised of 1% normal donkey serum (NDS) for 40 minutes. A rabbit anti-CD11b primary antibody was diluted in NDS and added to the slides for 60 minutes. After washing, slides were incubated for 40 minutes with a fluorescently tagged secondary antibody. The staining process was then repeated, on all slides, for the labeling of CD206. Following washes, DAPI (4’,6-diamidino-2-phenylindole) was used to label the nuclei of each section. Slides were mounted using a homemade anti-quenching mounting media containing Mowiol (Calbiochem #475904) and DABCO (Sigma #D2522) and imaged with a Zeiss Axio Slide Scanner. Antibodies are shown in Supplementary Table 4.

Statistical analysis

Correlation analyses were performed using the non-parametric Spearman rank correlation method (two-tailed, 95% confidence) using exact permutation P values when ranks were used, or Peason’s correlation test when both variables were continuous. Analysis of variance (ANOVA) was used for comparing statistical differences between multiple groups, followed by Dunn’s multiple comparison tests. Kruskal-Wallis ANOVA or Mann-Whitney tests were used. All statistical analyses were performed using GraphPad Prism (version 8.4.2 GraphPad Software, La Jolla California USA) and R software (URL http://www.R-project.org/). Correlation plots were created using the ‘Performance Analytics’ and ‘corrr’ plots, respectively, in R51–53.

Competing Interests statement

These authors declare no competing interests.

Author Contributions

MDF analyzed data composed figures and wrote the manuscript, BRV was the lead pathologist and LADM was the project veterinarian, contributed to study design, and writing of the IACUC and manuscript. CCM designed IHC panels and performed the IHC staining, GZ wrote the manuscript, KERL was a project veterinarian, made clinical assessments and collected samples, CJM processed and analyzed samples for RT-qPCR, contributed; EHH provided the mesoscale data, TPP stained the samples with GL and BMT. GL contributed to study design, provided administrative support, and aided with sample processing and archiving. NG contributed to study design, study coordination, sample processing, and SOP development. PKD processed and analyzed viral load data. CJR conceived and performed aerosol experiments. RPB contributed to study design, analysis of clinical and imaging results, and writing the manuscripts. NJM performed staining of BAL and contributed to writing the manuscript. TF contributed to the animal study design and planning and contributed to writing the manuscript. JR designed and supported the animal study and helped with the writing of the manuscript. MV conceived and supported the study, run experiments, analyzed data and wrote the manuscript.

Figure Legend

View this table:
Supplementary Table 1: Breakdown of routes of exposure and viral inoculation
View this table:
Supplementary Table 2: Virus loads in clinical specimens
View this table:
Supplementary Table 3: Rotation table derived from PCA
View this table:
Supplementary Table 4: Antibodies used in flow cytometry and immunofluorescent microscopy panels
Supplementary Figure 1.
Supplementary Figure 1. Kinetics of monocytes in Rhesus Macaques and African Green Monkeys.

Association between SARS-CoV-2 load in bronchial brushes and ranking of disease severity in AGM (A) and RM (B) (most severe case = 1; mildest case = 8). Changes in monocyte count in blood before and after infection in AGM (C) and RM (D). Proportion of total monocytes (HLA-DR+ CD16+ and/or CD14+ out of CD45+ leukocytes) in PBMCs from AGM (E) and RM (F) over time. (G) Changes in the absolute number of neutrophils in blood over time in all eight animals. Correlation between SARS-CoV-2 viral load in bronchial brushes and absolute count of monocytes in blood at one-week post- infection (1wk pi) in AGM (H) and RM (I). Percent of classical (CD14+ CD16–) monocytes (CD14– CD16+) out of total monocytes in PBMCs from AGM (J) or RM (K). Percent of non classical monocytes (CD14– CD16+) out of total monocytes in PBMCs in AGM (L) or RM (M). (N) Percent of intermediate monocytes (CD14+ CD16+) out of total monocytes in PBMCs from all eight animals over time.

Supplementary Figure 2.
Supplementary Figure 2. Chemokines promote cell migration of monocytes from the blood to the lung following infection.

Levels of chemokines in the peripheral blood over time, including (A) Eotaxin, (B) IP-10, (C) MCP-1 (CCL2), (D) MCP-4 (CCL13), (E) MDC (CCL22), (F) MIP-1a, (G), MIP-1b, and (H) TARC (CCL17). (I) PCA plot comparing baseline levels of chemokines between rhesus macaques and African Green monkeys. (J) TARC (CCL17) levels and (K) IP-10 levels in blood and SARS-CoV-2 viral load in bronchial brushes one week post-infection.

Supplementary Figure 3.
Supplementary Figure 3. Changes in the frequency of alveolar macrophages and infiltrates in BAL during infection by flow cytometric analysis.

(A) Left panel. Representative light scatter (forward scatter and side scatter) analysis of lymphocytes, macrophages, and polymorphonuclear cells (PMNC) in BAL. Cell populations are located according to their relative size (forward scatter) and granularity (side scatter). Right panels. Overlaid histograms showing fluorescent intensities of CD206 and HLA-DR of lymphocytes, macrophages, and PMNC as shown on the left panels. (B) Representative pseudocolor dot plot displaying alveolar macrophages, endothelial cells, monocytes, and/or dendritic cells distinguished by differential expression of CD163 and CD206. (C) Range of phenotypes of CD11b+ and CD11c+ myeloid cells in BAL and their change in proportion out of total HLA-DR+ cells over time. (D) tSNE plot displaying kinetics of CD86 expression on total HLA-DR+ cells after infection. Percent of CD16+ HLA-DR+ CD206- cells in BAL over time in AGM (E) and RM (F). (G) Representative gating strategy showing classification of CD45+HLA-DR+CD16+ myeloid cells which express either CD11c or CD11b. Top panel showing a sample with CD11c+ cells, and bottom panel showing a sample without CD11c+ cells used to help determine optimal gate position.

Supplementary Figure 4.
Supplementary Figure 4. Association of myeloid subsets in the BAL with IL-6 plasma levels.

Correlation plot between myeloid subsets in BAL and levels of IL-6 (pg/mL) in the blood 3 weeks post infection using Pearson’s method. Numbers inside boxes on the diagonal right half of the correlation plot represent the coefficient of correlation (r) and asterisks denote significance. One red asterisk corresponds with p <. 05, two red asterisks correspond with p < .01, and three red asterisks correspond with p < .001. Correlation plots in the bottom left display the actual data points.

Supplementary Figure 5.
Supplementary Figure 5. Lymphocyte response to SARS-CoV-2 in Rhesus Macaques and African Green Monkeys.

Percent of NKG2a+ on CD3- cells in BAL in AGM (A) and RM (B). Changes in lymphocyte count in blood before and after infection in AGM (C) and RM (D). (E) Median levels of IP-10, IFN-γ, IL-13, and IL-5 in plasma (pg/mL) from all eight animals over time. (F) Correlation between IL-4 levels in the plasma (pg/mL) and percent of PD-1+ Th2 cells in the blood at week 3 post infection. Proportion of PD-1+ CD8+ T cells in BAL from AGM (G) and RM (H) after infection.

Acknowledgments

We would like to thank Dr. Joseph (JC) Mudd and Dr. Cariappa Annaiah for critical reading of the manuscript and Amitinder (Miti) Kaur, Megan Varnado, Kaitlin Didier from the TNPRC Flow Cytometry Core for their help. We thank Angela Birnbaum for reviewing and optimizing all technical SOPs and overseeing the safety of this study.

We would also like to thank Fast Grant Funding for COVID-19 Science for partially funding this work and the NIH for supporting this work through the TNPRC base grant (P51 OD011104 59).

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Posted July 21, 2020.
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Cellular events of acute, resolving or progressive COVID-19 in SARS-CoV-2 infected non-human primates
MD Fahlberg, RV Blair, LA Doyle-Meyers, CC Midkiff, G Zenere, KE Russell-Lodrigue, CJ Monjure, EH Haupt, TP Penney, G Lehmicke, BM Threeton, N Golden, PK Datta, CJ Roy, RP Bohm, NJ Maness, T Fischer, J Rappaport, M Vaccari
bioRxiv 2020.07.21.213777; doi: https://doi.org/10.1101/2020.07.21.213777
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