Background
Human immunodeficiency virus peripheral neuropathy (HIV-PN) continues to be problematic among the HIV-infected population despite successful use of anti-retroviral therapy (ART) to reduce plasma viral loads and increase patient longevity [
1]. Distal sensory polyneuropathy (DSP), a common type of HIV-PN, consists of damage to the dorsal root ganglia (DRG) that is associated with loss of distal axons including small diameter unmyelinated nerve fibers that project to the skin and are measured by intraepidermal nerve fiber density (IENFD) [
2‐
4]. Axonal loss and damage to the DRG is thought to occur either via direct neurotoxicity of viral proteins or through indirect mechanisms from chronic immune activation. Application of HIV viral proteins, Tat, gp120, and Vpr in vitro has resulted in neurotoxicity of DRG neurons [
5‐
7]. However, neuronal damage can occur in humans with undetectable viral loads [
1], and viral expression in the DRG does not correlate with pathology severity [
8]. Instead, in the SIV rhesus macaque animal model, we found that monocyte traffic to the DRG is associated with DRG pathology, as well as a loss of IENFD [
8]. However, the underlying mechanism of neurodegeneration in vivo has yet to be fully elucidated.
Monocyte activation is a hallmark of HIV comorbidities among patients on ART [
9‐
11]. Monocyte and macrophage activation, often measured by plasma biomarkers such as sCD14 and sCD163, has been associated with an increase risk of cardiovascular disease, HIV-associated neurocognitive disorders (HAND), renal disease, and frailty [
12‐
15]. Additionally, a greater rate of monocyte egress from the bone marrow is associated with faster progression to AIDS in a CD8-depleted SIV-infected rhesus macaque model of HIV [
16]. Furthermore, the CD14+ CD16+ monocyte population expands during HIV infection [
17‐
19], and this expansion of activated monocytes is associated with HAND [
18,
20].
Cytokines have a large impact on the nervous system, in addition to regulating the immune response. DRG neurons express cytokine receptors on their surfaces, so they can appropriately respond to cytokines in their environment [
21,
22]. CCR5, the receptor for RANTES (Regulated on Activation, Normal T cell Expressed and Secreted)/CCL5, can be expressed on DRG neuronal cell bodies. gp120 can bind to CCR5 resulting in neuronal excitation [
23]. CCR2, another chemokine receptor expressed on DRG neurons, can also facilitate neuronal excitation when it binds to monocyte-chemoattractant-1 (MCP-1/CCL2) [
24]. Various rodent models of neuropathic pain have demonstrated that blocking the MCP-1-CCR2 interaction via neutralizing antibodies or gene knockout can block pain sensation [
25‐
29]. Neuronal cell bodies in the DRG can also upregulate inflammatory cytokines following peripheral nerve injury [
30‐
32]. Thus, cytokines can transmit pain signals from the periphery to the central nervous system (CNS) via interactions with cytokine receptors on DRG neurons [
28,
33]. Additionally, MCP-1 and RANTES might increase recruitment of monocytes to the DRG causing further neuronal damage and activation [
21,
34,
35].
Other signaling pathways besides CCR2 and CCR5 are likely involved in neuronal damage during SIV-DSP. One potential signaling protein of interest is CD137, which is a member of TNF superfamily that can be expressed on T cells, monocytes, and other immune cells, as well as endothelial cells [
36,
37]. CD137 cross-linking on monocytes induces activation and production of pro-inflammatory cytokines [
38]. CD137 expression on endothelial cells facilitates migration of monocytes out of blood vessels and into tissues [
37,
39]. Additionally, CD137 reverse signaling is involved in myelopoiesis [
40,
41]. Elevated sCD137 in plasma, a splice variant of CD137, has been associated with several inflammation-linked diseases [
42‐
44], but its role in monocyte activation during HIV infection has not been studied.
We used a CD8-depleted, SIV-infected macaque model to recapitulate HIV-DSP in humans, where animals show a loss of IENFD and DRG pathology [
45‐
47]. We have previously demonstrated an influx of activated MAC387+ macrophages to the DRG as well as an increase in CD163+ macrophages. Importantly, we found that increased cell traffic was associated with severe DRG pathology and a greater loss of IENFD [
8]. This study sought to investigate the role of monocyte activation in HIV-DSP, as well as identify cytokines that are associated with monocyte activation and neuronal loss in plasma and in DRG tissue.
Methods
Ethical statement
All animals used in this study were handled in strict accordance with American Association for Accreditation of Laboratory Animal Care with the approval of the Institutional Animal Care and Use Committee of Harvard University and the Institutional Animal Care and Use Committee of Tulane University.
Animals, viral infection, and CD8 lymphocyte depletion
Fifteen rhesus macaques (Macaca mulatta) were utilized in this study. Eleven animals were inoculated intravenously with SIVmac251 (a generous gift from Dr. Ronald Desrosiers, University of Miami). Four uninfected rhesus macaques served as uninfected controls. All infected animals were administered 10 mg/kg of anti-CD8 antibody subcutaneously at day 6 after infection and 5 mg/kg intravenously at days 8 and 12 after infection in order to achieve rapid progression to AIDS. The human anti-CD8 antibody was provided by the NIH Non-human Primate Reagent Resource (RR016001, AI040101). SIV-infected animals were sacrificed at the onset of terminal AIDS. The development of simian AIDS was determined post-mortem by the presence of Pneumocystis carinii-associated interstitial pneumonia, Mycobacterium avium-associated granulomatous enteritis, hepatitis, lymphadenitis, and/or adenovirus infection of surface enterocytes in both small and large intestine. Animals were housed at either the New England Primate Research Center (NEPRC; Southborough, MA) or Tulane University’s National Primate Research Center (TNPRC; Covington, LA) in strict accordance with standards of the American Association for Accreditation of Laboratory Animal Care.
Necropsy and histopathology
Animals were necropsied immediately following death, and representative sections of all major organs were collected, fixed in 10 % neutral-buffered formalin (NBF), embedded in paraffin, and sectioned at 5 μm. After deparaffinization in xylene, the tissues were hydrated in graded alcohols, counterstained with Harris hematoxylin solution (Sigma-Aldrich) for two minutes, and rinsed with running water. The slides were then dipped sequentially in acid alcohol (90 % methanol, 5 % sulfuric acid, 5 % acetic acid; Sigma-Aldrich) and ammonia water (15–20 drops ammonium hydroxide in 250 ml water; Sigma-Aldrich), rinsing with running water after each, followed by 80 % alcohol for 2 min and eosin (Sigma-Aldrich) for 2 min. Tissue sections were then rinsed in graded alcohols and dehydrated with xylene and mounted with VectaMount (Vector).
Histopathologic analysis of DRG morphology
H and Estained sections of DRG were evaluated blindly for histopathologic lesions by a board-certified veterinary anatomic pathologist (ADM) and scored based on the presence and severity of infiltrating mononuclear cells, neuronophagia, and Nageotte nodules as previously described [
8,
46]. Overall pathology was scored on a previously validated [
8] scale of 1–3 at increments of 0.5 via the following criteria: (1) Mild: scattered infiltrating mononuclear cells with rare evidence of neuronophagia and/or neuronal loss, (2) Moderate: increased numbers of infiltrating mononuclear cells with occasional neuronophagia and/or neuronal loss, and (3) Severe: abundant infiltrating mononuclear cells, frequent neuronophagia and neuronal loss were all present [
8,
46,
48].
Immunohistochemistry
DRG sections were deparaffinized with xylene and hydrated in a series of graded alcohols. Sections were stained with antibodies against MAC387 (clone M0747; Dako), CCR5 (rabbit polyclonal; Novus Biologicals) or CCR2 (clone 7A7; Abcam). Frozen DRG sections were used for CD137 staining (clone BBK-2). Sections were counterstained with hematoxylin, dehydrated, and mounted using VectaMount permanent mounting medium (Vector Labs). Tissues were visualized using a Zeiss Axio Imager M1 microscope (Carl Zeiss MicroImaging). Quantification of the absolute number and percent of positive satellite cells were performed as previously described [
8]. For each animal, eight non-overlapping fields of view at ×200 magnification were quantified by manually counting the number of positive cells in the field and dividing by the total area of DRG tissue. The average number of positive cells per square millimeter was used.
Skin punch and IENFD measurement
Skin punch biopsy specimens with IENF were performed in all SIV+ animals. Skin punches (3 mm) were taken serially near the sural innervation site just distal to the lateral malleolus. Biopsy specimens were taken for each animal at pre-infection, several time points during infection, and at necropsy. Biopsy specimens were fixed in Zamboni’s fixative and processed for dividing into sections. Sections (50-μm thick) of serial punch skin biopsy specimens were stained with anti-PGP 9.5, a panaxonal marker (1:10,000 dilution; ABD Serotec). Nerve fiber length/volume of epidermis (IENFD) was quantified using computer software (Space balls program; Microbrightfield Bioscience) as previously described [
8,
49].
BrdU administration
A 30 mg/mL stock of solution was prepared by adding 5-bromo-2-deoxyuridine (BrdU; Sigma-Aldrich) to 1× phosphate-buffered saline (without Ca
2+ and Mg
2+) and heated to 60 °C in a water bath, as previously described [
8,
16,
50]. BrdU was administered as a slow bolus i.v. injection at a dose of 60 mg BrdU/kg body weight. BrdU was administered at 8 and 21 days post-infection (DPI) in animals A01-A07. Additionally, animals A04-A11 received BrdU 42, 63 DPI, and 24 h prior to necropsy.
Flow cytometry
Flow cytometric analyses were performed with 100 μl aliquots of EDTA-coagulated whole blood. Erythrocytes were lysed using ImmunoPrep Reagent System (Beckman Coulter), washed twice with PBS containing 2 % FBS, and then incubated for 15 min at room temperature with fluorochrome-conjugated surface antibodies including anti-HLA-DR-PerCp-Cy5.5 (clone L243), anti-CD16-PE-Cy7 (clone 3G8), anti-CD3-APC (clone SP34-2), CD8-APC (clone RPA-T8), anti-CD20-APC (clone 2H7), and anti-CD14-Pacific blue (clone M5E2). For intracellular staining, cells were fixed and permeabilized with BD Cytofix/Cytoperm™ buffer (BD Biosciences) for 30 min at room temperature. Cells were again washed and incubated with BD Cytoperm Plus™ buffer for 10 min on ice, then washed and incubated with DNase (30 mg) for 1 h at 37 °C, and washed and then stained for intracellular antigen with anti-BrdU-FITC (clone 3D4; BD Biosciences) and anti-Ki-67-PE (clone B56; BD Biosciences) for 20 min at room temperature. Samples were acquired on a BD FACS Aria (BD Biosciences) and analyzed with Tree Star Flow Jo version 9.6. Identification and quantitation of BrdU+ monocytes and CD14+ CD16+ monocytes was performed as previously described [
16].
Preparation of DRG lysate
Frozen lumbar DRG was mechanically homogenized in Tissue Extraction Reagent I (Invitrogen, Waltham, MA) containing 1× protease inhibitor (Sigma-Aldrich). For every 1 g of tissue, 10 mL of lysis buffer was used. Lysate was centrifuged and supernatant containing protein was stored at −80 °C. Protein was quantified using a BCA protein assay kit (Thermo Scientific) according to the manufacturer’s instructions.
ELISAs
sCD14 and RANTES were quantified in plasma (diluted 1:200 and 1:4; respectively) using ELISA kits (R&D Systems). sCD163 was quantified in plasma (diluted 1:500) using an ELISA kit (Trillium Diagnostics). All ELISAs were carried out according to the manufacturer’s instructions and as previously described [
16].
Luminex multiplex assays
RANTES, MCP-1, and sCD137 were quantified in DRG tissue lysates, and MCP-1 and sCD137 were quantified in plasma using Multiplex Luminex Technology (EMD Millipore). Non-human Primate Cytokine/Chemokine Panels 1 and 2 were used according to the manufacturer’s instructions with the following modifications. For DRG tissue lysate protein quantification, 10 μg of protein (in 25 μl of lysis buffer) from each sample was loaded onto the plate. Tissue lysis buffer was used as the matrix for dilution of standards and quality control samples. For the plasma sample analysis, plasma samples were diluted two-fold in the assay buffer provided. The provided serum matrix was used for dilution of standards and quality control samples. All samples were performed in duplicate, and plates were incubated overnight at 4 °C on a rocker. Samples were analyzed using MAGPIX System (EMD Millipore).
Statistical analysis
All statistical analysis was performed using Prism Software (Version 5.0d). A Wilcoxon matched-pairs signed-rank test was used to determine the increase in markers from pre-infection to necropsy. A Mann-Whitney U test was used to detect variation between infected and uninfected samples. ANOVA was used to detect variance among different pathology groups followed by a Dunn’s post-test if the ANOVA was significant. Non-parametric Spearman correlation was used for all correlations. A p value of <0.05 was considered significant for all tests performed.
Discussion
HIV-PN continues to be a major co-morbidity of HIV infection despite reduction of plasma viral load. Currently, there is no successful treatment for HIV-PN, and thus, understanding the underlying mechanism of neuronal damage is of utmost importance to improve patient quality of life. There is increasing evidence that chronic monocyte/macrophage activation plays a key role in the pathogenesis of HIV/SIV-PN. Animal models of HIV-PN and studies in humans have shown an increase in myeloid cell activation in the DRG and spinal column, as well as a loss of IENFD [
3,
8,
53‐
55]. We have previously demonstrated that increased BrdU+ monocyte traffic to the DRG is associated with severe DRG pathology [
8].
Numerous studies have examined the expansion of the CD14+ CD16+ monocyte population during HIV/SIV infection and the link to inflammatory co-morbidities [
16,
56,
57]. We have also shown that the rate of monocyte egress from the bone marrow, measured by BrdU pulse labeling, is correlated to faster disease progression to AIDS [
16]. To investigate the systemic inflammation that is causing neuronal damage, both in the DRG and in the extremities, we examined five soluble proteins in plasma, which are associated with monocyte activation and traffic. Here, we found that plasma sCD14 and MCP-1 were correlated to the number of BrdU+ monocytes in blood. We also found that sCD14, RANTES, and MCP-1 were correlated to the percent of CD14+ CD16+ monocytes out of total blood monocytes. CD14+ CD16+ monocytes are an activated population of monocytes that highly express CCR2, the receptor for MCP-1, and CD163 [
58,
59]. Elevated levels of RANTES and MCP-1 in plasma were associated with moderate or severe DRG pathology, compared to mild pathology. These findings confirm these soluble factors in plasma are associated with monocyte activation and traffic during SIV-PN.
Because the blood-nerve barrier is more promiscuous (or leakier) than the blood-brain barrier [
60], we assumed that neurons were exposed to all proteins found in plasma. Several inflammatory cytokines have been found to be neurotoxic in vitro [
61,
62], but this direct causation is difficult to prove in vivo
. We found a significant inverse correlation between sCD163 and IENFD. Thus, sCD163 may be useful as a plasma biomarker of IENFD loss in HIV+ patients. While sCD163 and sCD14 are both markers of monocyte activation, they are shed by different mechanisms. CD163 is highly expressed on M2-polarized macrophages and CD14+ CD16+ monocytes, while CD14 is present on all populations of monocytes and is shed in the setting of non-specific activation [
63] and CD163 is shed due to cell-surface TLR activation [
64]. Plasma RANTES/CCL5 also correlated to a reduction in IENFD. Other studies have demonstrated that the supernatant of macrophages exposed to gp120, presumably containing proteins such as sCD163 and RANTES, is capable of damaging neurons in vitro [
61]. Even though sCD163 and RANTES strongly correlated to a reduction of IENFD, the dying back of axons is likely caused by several signals.
Here, we found a link between the severity of DRG pathology and a greater loss of small nerve fiber density in the footpad. It is unknown if a loss of IENFD comes before damage to the DRG or vice versa. One study suggested that damage to the DRG preceded altered functional activity of nerve fibers in the periphery [
65]. However, we have observed an early loss of IENFD (as early as 8 DPI) and minimal DRG pathology in animals sacrificed at 21 DPI (data not shown). Regardless, we found an association between pathology at the DRG and in the footpad, suggesting a relay of signals from one region to the other, or perhaps systemic neuroinflammatory/neurotoxic proteins that facilitate damage to both regions simultaneously.
To investigate the local signals in the DRG responsible for monocyte traffic, we analyzed DRG tissue homogenate using a multiplex assay that allowed for quantification of many proteins with a small amount of tissue homogenate available. The limitation of this method is that it is unknown which cell types are producing the proteins that were detected. Endothelial cells, neurons, Schwann cells, and immune cells (including macrophages and T cells) in the DRG are all capable of secreting cytokines and chemokines. However, this method still affords us the opportunity to investigate the local signals responsible for monocyte traffic and neuronal damage at the DRG. We found that MCP-1 in DRG tissue was significantly increased in DRG from SIV+ animals compared to uninfected control tissue. Because MCP-1 is a potent monocyte chemoattractant and is produced by activated macrophages, it is likely that MCP-1 is partially responsible for increased monocyte traffic to the DRG.
In addition to the proteins we reported on in detail here, we also investigated other known monocyte chemoattractants, both in the DRG and in plasma. We did not find a significant increase in MIP-1α, MIP-1β, and MIP-3α in plasma nor were these proteins elevated in SIV+ DRG tissue lysate. In fact, these proteins were below detection level for many of the DRG samples tested. However, monocyte activation and traffic is a complex process, likely to be controlled by several signaling molecules that were not included on the two cytokine/chemokine panels we utilized.
sCD137/CD137 (formally called 4-1BB or tumor necrosis factor receptor superfamily member 9 (TNFRSF9)) has not been extensively studied in the context of monocyte activation during HIV infection, although its known functions in other diseases are relevant to HIV pathogenesis. Here, CD137 was found to potentially play a role in SIV-DSP pathogenesis. The soluble form of CD137 (sCD137) is generated by alternative splicing and was found to significantly increase in plasma from pre-infection to terminal AIDS. Additionally, sCD137 was only detectable in DRG lysate with severe pathology, and it correlated with the number of MAC387+ cells in DRG tissue. CD137 is expressed on a wide range of cell types, although most of the research on this protein focuses on T cell activation [
36‐
38]. However, CD137 signaling has been shown to play a role in myelopoiesis, monocyte activation, and monocyte extravasation into inflamed tissue [
37,
38,
41]. The known roles of CD137 in regard to monocyte activity are also highly deregulated during HIV/SIV infection. Additional research needs to be conducted in order to further define the role of CD137 signaling in HIV/SIV disease progression. CD137 activation, through the use of agnostic monoclonal antibodies, has proven to have potential for cancer treatment by stimulating the immune system to target cancer cells [
66]. Blocking CD137 may ameliorate chronic immune activation seen during HIV/SIV infection.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
THB and JRL conceived and designed the experiments. JRL, JAR, and MJP performed the experiments. THB, ADM and JRL analyzed the data. THB and JRL wrote the paper. MJP and ADM carried out the paper revision. All authors read and approved the final manuscript.