Exogenous pericyte delivery protects the mouse kidney from chronic ischemic injury

Abstract

Pericytes are considered reparative mesenchymal stem cell-like cells, but their ability to ameliorate chronic ischemic kidney injury is unknown. We hypothesized that pericytes would exhibit renoprotective effects in murine renal artery stenosis (RAS). Porcine kidney-derived pericytes (5 × 10⁵) or vehicle were injected into the carotid artery 2 wk after the induction of unilateral RAS in mice. The stenotic kidney glomerular filtration rate and tissue oxygenation were measured 2 wk later using magnetic resonance imaging. We subsequently compared kidney oxidative stress, inflammation, apoptosis, fibrosis, and systemic levels of oxidative and inflammatory cytokines. Treatment of xenogeneic pericytes ameliorated the RAS-induced loss of perfusion, glomerular filtration rate, and atrophy in stenotic kidneys and restored cortical and medullary oxygenation but did not blunt hypertension. Ex vivo, pericytes injection partially mitigated RAS-induced renal inflammation, fibrosis, oxidative stress, apoptosis, and senescence. Furthermore, coculture with pericytes in vitro protected pig kidney-1 tubular cells from injury. In conclusion, exogenous delivery of renal pericytes protects the poststenotic mouse kidney from ischemic injury, underscoring the therapeutic potential role of pericytes in subjects with ischemic kidney disease.

NEW & NOTEWORTHY Our study demonstrates a novel pericyte-based therapy for the injured kidney. The beneficial effect of pericyte delivery appears to be mediated by ameliorating oxidative stress, inflammation, cellular apoptosis, and senescence in the stenotic kidney and improved tissue hypoxia, vascular loss, fibrosis, and tubular atrophy. Our data may form the basis for pericyte-based therapy, and additional research studies are needed to gain further insight into their role in improving renal function.

INTRODUCTION

Renal artery stenosis (RAS) is increasingly recognized as a common cause of ischemic kidney disease and renal dysfunction. What is worse, the presence of RAS is an independent risk factor for death (1). Hemodynamically significant RAS can be detected in 6.8% of individuals over 65 yr old and in up to 40% of individuals with vascular disease (2,3), implying that a large segment of the population may require medical attention. However, a recent clinical trial showed that renal revascularization that abolishes the stenosis fails to improve renal function, decrease renal or cardiovascular events, or increase patient survival (4). As an alternative to the present limited therapy, stem cell therapy is deemed promising for the restoration of kidney function and prevention of progressive renal injury (5).

Pericytes are mural cells encircling the microvascular endothelium of capillaries, terminal arterioles, and postcapillary venules (6). They are found in most tissues at a ratio of 1:1, 1:2.5, and 1:100 of pericytes to endothelial cells in the retina, kidney, and skeletal muscle, respectively (7, 8). After an injurious insult, renal pericytes may detach from the vasculature, driving microvascular rarefaction and subsequent hypoxia associated with chronic kidney disease (9). In addition, they have emerged as major contributors to activated matrix-depositing stromal cell populations seen in progressive renal fibrosis (8). Intriguingly, pericytes have also been suggested to locate within a mesenchymal stem/stromal cell (MSC) niche, express MSC markers (10), and differentiate into chondrocyte, adipocyte, and osteocyte lineages (10, 11), indicating that they may harbor MSC-like properties. Indeed, exogenous delivery of pericytes ameliorates ischemic injury in the heart (12), skeletal muscles (13), and retinal vasculopathy (14). Thus, owing to their wide distribution in the microvasculature, pericytes might represent a promising and attractive source of precursor cells for regenerative medicine. However, whether exogenously delivered pericytes can attenuate chronic ischemic kidney injury remains unknown.

To address this question, we investigated the therapeutic potential of primary swine kidney-derived pericytes in a chronic ischemic renal injury model in mice. Transplantation of pericytes improved not only renal blood flow but also kidney function. Major repair mechanisms included reduction of fibrosis, inhibition of chronic inflammation, promotion of angiogenesis, and attenuation of senescence of tubular cells.

METHODS

Study Animals

The Mayo Clinic Institutional Animal Care and Use Committee approved this study (protocol no. A00004509-19). Seventy-two male 129S1 mice and two healthy farm pigs were used. All animal procedures were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

Male 129S1 mice (RRID:MGI:2671655, Jackson Laboratory, 11 wk of age) were randomly divided into sham, sham + pericyte-treated, RAS + vehicle-treated, and RAS + pericyte-treated groups (n = 8/group) and studied for 4 wk. To evaluate whether the effects of pericytes were shared among other reparative cells, for limited comparison purposes, an additional group of RAS mice received porcine renal scattered tubular-like cells (STCs; n = 8) and were also observed for 4 wk. We have previously shown that porcine STCs protected against kidney tubular cell injury in murine stenotic kidneys with RAS as well as in vitro, and STCs were harvested from healthy pig kidneys as previously described (15).

RAS was induced as previously described (16). After a right flank incision (∼1.5 cm), we surgically placed a 0.15-mm-diameter arterial cuff around the right renal artery and secured it by two 10-0 nylon circumferential sutures (Surgical Specialties, Reading, PA); sham surgeries excluded cuff placement (17). This approach leads to a fall in renal volume, blood flow, and function. Blood pressure was measured at baseline and 2, 3, and 4 wk after surgery by tail cuff (Kent Scientific, Torrington, CT).

After 2 wk, the carotid artery was cannulated and 200-µL PBS (RAS + vehicle), porcine pericytes (5 × 10⁵ cells in 200-µL PBS), or porcine STCs (5 × 10⁵ cells in 200 µL PBS) were slowly injected caudally into the aorta. All pericytes had been prelabeled with cell trace far red (C34564, ThermoFisher). To trace pericyte distribution by flow cytometry, in some sham + pericyte- and RAS + pericyte-treated mice, we also collected visceral solid organs, including the heart, lung, liver, spleen, and both left and right kidneys (n = 3 each), 48 h after injection. Two weeks after the injection, mice were scanned with magnetic resonance imaging (MRI) and subsequently euthanized with CO₂. Kidneys and blood samples were collected for ex vivo experiments.

Cell Culture

Pericytes were identified and isolated from four freshly harvested healthy pig kidneys (6 mo old) as previously described (18) with few modifications. First, pericytes were localized and visualized within the excised pig kidney on 5-µm slices by immunofluorescence. Then, 3- to 5-g pig kidneys with both the cortex and medulla were resected and washed twice with PBS. Kidney tissue was then diced and digested with 2 mg/mL collagenase for 1 h and filtered by a 60-mesh (250-μm) steel sieve to remove undigested tissue fragments. The cellular components were then passed through a 100-μm cell strainer followed by the addition of a prepared pericyte growth medium at 37°C in a humidified atmosphere with 5% CO₂. For in vitro experiments, pig pericytes were cultured in pericyte growth medium (100 mL), which consisted of 10-mL FBS (Cat. No. F2442, Sigma), 1-mL penicillin-streptomycin solution (Cat. No. P4333, Sigma), 1-mL PC Growth Supplement 100 (Cat. No. 1252, Science), and 88-mL low-glucose DMEM (Cat. No. 11885-084, Life Technologies). Pigment epithelium-derived factor (100 nM, Cat. No. SRP4988, Sigma) was added to the medium on day 2 (19). The medium was replenished every 2–3 days to remove nonadherent cells, and passages with 1:2 were performed once cells reached 70%–80% confluency. After three passages, cells were harvested for phenotype analysis with immunofluorescence for neural/glial antigen-2 (NG-2; Cat. No. NBP1-21361, Novusbio, 1:100), α-smooth muscle antigen (α-SMA; Cat. No. ab7817, Abcam, 1:200), CD31 (Cat. No. MCA1747, Bio-Rad, 1:50), and platelet-derived growth factor (PDGF) receptor (PDGFR)-β (Cat. No. 323606, BioLegend, 1:100).

Renal Hemodynamics and Oxygenation

Twelve mice in each group (and 8 RAS + STC-treated mice) were scanned with a 16.4-T MRI scanner, and renal volume and hemodynamics were assessed as previously described (17). Renal volume was quantified from images acquired using respiration-gated three-dimensional fast imaging. Renal perfusion was measured with arterial spin labeling and quantified from the flow-sensitive alternating inversion-recovery sequence with rapid acquisition with relaxation enhancement images. Renal perfusion was measured in RAS + STC-treated mice as well. Renal oxygenation was assessed with blood oxygen level-dependent MRI. Eight images were reconstructed after zero filling the k-space data to 256 × 256. T2* was quantified by pixel-wise monoexponential fitting on the averaged magnitude of all eight images over echo times. R2* (1/T2*) was used as an index of blood oxygenation level. Renal gene expression of hypoxia-related molecules such as hypoxia-induced factor (HIF)-1α, vascular endothelial growth factor (VEGF), and VEGF receptor (VEGFR) was also assessed by quantitative PCR.

Renal Function

Single-kidney glomerular filtration rate (GFR) was evaluated by Dynamic Contrast-Enhanced MRI (20). Renal function was also assessed by the levels of plasma creatinine, blood urea nitrogen, and urine protein using commercial kits (DetectX kits, Arbor Assays, Ann Arbor, MI). Briefly, 25 μL of standards or samples were pipetted into a microliter plate, and the color-generating reaction was initiated with the reagent. The concentration of creatinine was calculated using the change (Δ) of the optical density readings at 30 min compared with the standard curve.

Renal Injury

Renal tissue was fixed in 10% neutral buffered formalin, dehydrated, and embedded in paraffin per standard techniques. Tubular injury was scored in a blinded fashion in sections stained with periodic acid-Schiff, as previously described (21). Absence of the brush border, dilation, atrophy, cast formation, cell detachment, or thickening of the tubular basement membrane were scored as 1–5, where 0 = normal tubules, 1 ≤ 10% injured tubules, 2 = 10%–25% of injured tubules, 3 = 26%–50% of injured tubules, 4 = 51%–75% of injured tubules, 5 ≥ 75% of injured tubules. Tubulointerstitial fibrosis was assessed in midhilar cross sections stained with Masson’s trichrome staining, semiautomatically quantified in 15–20 fields. Results were averaged from all fields and expressed as percent area staining. In addition, expression of transforming growth factor (TGF)-β was evaluated by Western blot analysis in each kidney sample using specific polyclonal antibodies (Santa Cruz Biotechnology, 1:200) (22).

Inflammation and Rejection

Renal inflammation was evaluated by immunohistochemistry staining with antibodies against macrophages (F4/80, Cat. No. ab100790, Abcam), and rejection by colocalization of pericytes with CD3 (Cat. No. ab16669, Abcam) T lymphocytes. Positive cells were manually counted in 20 fields of random glomerular or cortical fields in each sample in a blinded manner. In addition, renal expression of tumor necrosis factor (TNF)-α, monocyte chemoattractant protein (MCP)-1, interferon (INF)-γ, and NF-κB was quantified by quantitative PCR. Systemic inflammation was assessed by inflammatory factor levels, such as interleukin (IL)-1α, IL-1β, IL-6, IL-10, and TNF-α, by Luminex (PCTTMAG-23K, Millipore).

Oxidative Stress, Apoptosis, and Senescence

Renal redox status was evaluated by dihydroethidium (DHE; Cat. No. D11347, Invitrogen) staining for detection of in situ production of superoxide anion in 30-µm frozen sections (23). Apoptosis was assessed in 5-μm midhilar kidney cross sections stained with caspase-3 and TUNEL (Cat. No. G3250, Promega), with positive cells manually counted in 15–20 fields under fluorescence microscopy (ZEN 2012 Blue Edition, Carl Zeiss) (16). Kidney tissue senescence was assessed by staining with spider senescence-associated β-galactosidase (SA-β-gal; SPiDER-βGal, SKU: SG04, Dojindo Molecular Technologies).

Cell Injury In Vitro

To evaluate the direct reparative potency of pericytes, coculture experiments were performed using endothelial and tubular cell lines. Briefly, human umbilical vein endothelial cells (HUVECs; Cat. No. 200k-05f, Cell Applications, San Diego, CA) were grown in endothelial cell growth medium (EGM-Plus Endothelial Cell Growth Media-Plus Bulletkit Medium, Cat. No. CC-5035, Lonza, Cohasset, MN), seeded at 5 × 10⁵ cells/well in a Transwell plate (Cat. No. 2944-076, VWR, Radnor, PA), and divided into four groups. Cells in groups 1 and 2 were cultured under normal conditions, whereas cells in groups 3 and 4 were coincubated with TNF-α (10 ng/mL, R&D Systems) and TGF-β1 (5 ng/mL, R&D Systems) for 3 days to induce endothelial cellular injury (24). This medium was then replaced with a fresh growth medium. In addition, groups 2 and 4 were subsequently cocultured with pericytes (2.5 × 10⁵ cells/well insert) for another 24 h. Afterward, all cells (n = 4 samples/group) were lysed and prepared for quantitative PCR.

In addition, a porcine kidney cell line (LLC-PK1, 5 × 10⁴ cells, CL-101, American Type Culture Collection) seeded on 24-well plates (lower chamber) were cocultured in a Transwell system with or without TNF-α (10 ng/mL) and antimycin-A (10 µmol/L, Sigma) for 24 h, to induce tubular cell injury (25). Pericytes (5 × 10⁴) were added to the upper chamber for 24 h. Lactate dehydrogenase (LDH) levels in the supernatants of the lower chamber released by injured PK1 cells were measured by a colorimetric kit (Cat. No. MAK066, Sigma).

Statistical Analysis

Statistical analysis was performed using JMP 14.0-Pro (Statistical Analysis System, Cary, NC, RRID:SCR_008567). Normally distributed data are expressed as means ± SD ANOVA and Student’s t tests were used to evaluate statistically significant differences among the groups. Non-normally distributed data are presented as medians (ranges), and a nonparametric comparison was performed (Wilcoxon, Kruskal-Wallis). P < 0.05 was considered as statistically significant.

RESULTS

Characterization of Pig Pericytes and Distribution

In pig kidneys, pericytes were localized around small vessels, as indicated by immunofluorescent NG-2⁺ cells outlining the shape of vessels. Pericytes were surrounded by α-SMA-expressing cells (Fig. 1A). In vitro, primary pericytes isolated from pig kidneys were characterized as PDGFR-β⁺, NG-2⁺, α-SMA⁻, and CD31⁻ cells (Fig. 1B). Two weeks after injection, we identified cell trace far red-stained pericytes in the kidneys, lung, and liver in both sham and RAS mice (Fig. 2A). Importantly, CD3⁺ lymphocytes in the kidney were not adjacent to pericytes, arguing against immune rejection (Fig. 2B). In sham mice, the two kidneys and liver had comparable numbers of pericytes, whereas the retention of pericytes in the lung was much higher. In RAS mice, the right kidney, liver, and lung had significantly higher pericyte retention than in sham mice, with no difference in the left kidney among the groups (Fig. 2C).

Figure 1.

A: pericytes are located in small vessels, surrounded by α-smooth muscle actin (α-SMA)⁺ cells. B: characterization of pig pericytes in vitro, positive for neural/glial antigen-2 (NG-2) and platelet-derived growth factor receptor (PDGFR)-β and negative for CD31 and α-SMA. n = 3.

Figure 2.

Distribution of pericyte after injection in mice with renal artery stenosis (RAS). A: 2 wk after pericyte injection, cell trace far red-stained pericytes were detected in the kidneys, lung, and liver in both sham and RAS mice. B: CD3⁺ lymphocytes did not cluster around kidney pericytes, arguing against immune rejection. C: in sham mice, flow cytometry revealed comparable numbers of pericytes in both kidneys and the liver; the retention of pericytes in the lung was much higher. In RAS mice, the right (R; stenotic) kidney, liver, and lung had significantly higher pericyte retention than in sham mice. Pericytes in the left (L) kidney were comparable among the groups. *P < 0.05 vs. the left kidney; †P < 0.05 vs. the right kidney; ‡P < 0.05 vs. the liver. NG-2, neural/glial antigen-2; PDGFR-β, platelet-derived growth factor receptor-β. n = 3. t test.

Pericytes Decreased Stenotic Kidney Tubular Injury and Fibrosis

Periodic acid-Schiff staining indicated minimal tubular injury in the sham and sham + pericyte-treated groups. RAS induced significant tubular atrophy, cast formation, and tubular brush border loss, which pericyte administration decreased. Trichrome and Sirius red staining detected sporadic focal interstitial fibrosis in stenotic RAS kidneys, which pericytes ameliorated; none was observed in sham and sham + pericyte-treated groups (Fig. 3A). Sham- and sham + pericyte-treated groups had comparable gene expression of TGF-β and plasminogen activator inhibitor-1, which were significantly upregulated in the RAS group but decreased in the RAS + pericyte-treated group, as did protein expression of TGF-β (Fig. 3, B and C).

Figure 3.

A: periodic acid-Schiff (PAS) staining detected minimal tubular injury in sham- and sham + pericyte-treated mice. Renal artery stenosis (RAS) induced significant tubular atrophy, cast formation, and brush border loss, which pericyte administration decreased. Trichrome and Sirius red staining identified sporadic focal interstitial fibrosis in stenotic kidneys, which pericytes ameliorated, and none in the sham- and sham + pericyte-treated groups. n = 8. t test. B: sham- and sham + pericyte-treated groups had comparable expression of transforming growth factor (TGF)-β and plasminogen activator inhibitor-1 (PAI-1) genes, which were significantly upregulated in the RAS group and decreased in the RAS + pericyte-treated group. C: sham- and sham + pericyte-treated groups had comparable expression of TGF-β protein, which was significantly upregulated in the RAS group but decreased in the RAS + pericyte-treated group. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. RAS. n = 4. t test.

Pericytes Ameliorated Systemic and Renal Inflammation

Superoxide anion production in situ as per DHE staining was elevated in RAS compared with Sham groups and partly decreased in the RAS + pericyte-treated group. Of note, sham-, sham + pericyte-treated, and RAS + pericyte-treated groups had similar numbers of CD3⁺ T cells, whereas RAS induced a significantly greater number of CD3⁺ T cells in the stenotic kidney. Similarly, RAS also increased the numbers of F4/80⁺ macrophages, which pericytes markedly decreased (Fig. 4A). Renal protein expression of MCP-1 and TNF-α was upregulated in RAS mice but decreased in RAS + pericyte-treated mice (Fig. 4B). Systemically, inflammatory markers such as IL-1α, IL-1β, IL-6, MCP-1, and TNF-α were greatly induced in the RAS group, but pericyte injection markedly decreased their levels (Table 1). IL-10 levels were unchanged. In vitro, we observed increased LDH release from injured PK-1 cells, which was lowered but not normalized by pericyte coculture (Fig. 4C).

Figure 4.

A: dihydroethidium (DHE) staining suggested that superoxide anion production in situ was elevated in the renal artery stenosis (RAS) group compared with the sham and sham + pericyte-treated groups and improved in the RAS + pericyte-treated group. Sham-, sham + pericyte-treated, and RAS + pericyte-treated groups had similar CD3⁺ T cell numbers, whereas RAS increased them in the stenotic kidney. RAS also increased F4/80⁺ macrophage infiltration, which pericytes decreased. n = 8. t test. B: renal gene expression of monocyte chemoattractant protein (MCP)-1 and tumor necrosis factor (TNF)-α was upregulated in RAS mice but markedly decreased in RAS + pericyte-treated mice. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. RAS. n = 5. t test. C: cell coculture found significant lactate dehydrogenase (LDH) release from injured PK-1 cells, which was significantly decreased by pericyte coculture. n = 5. t test. *P < 0.05 vs. PK-1; †P < 0.05 vs. PK-1 + pericytes; ‡P < 0.05 vs. PK-1 + injury.

Table 1.

Systemic characteristics and renal function in mice with RAS

	Sham	Sham + Pericytes	RAS	RAS + Pericytes
Body weight, g	24.6 ± 1.1	23.2 ± 2.4	19.2 ± 2.9*†	26.5 ± 1.7†‡
STK weight, mg	207 ± 29	184 ± 7	59 ± 4*†	135 ± 51‡
STK-to-contralateral kidney weight ratio	1.02 ± 0.02	0.97 ± 0.03	0.29 ± 0.03*†	0.78 ± 0.34‡
Serum creatinine, mg/dL	0.17 ± 0.05	0.22 ± 0.09	0.23 ± 0.02*	0.25 ± 0.04*
Blood urea nitrogen, mg/dL	22.7 ± 5.6	52.9 ± 9.4*	50.4 ± 4.7*	56.9 ± 12*
Urine protein, mg/dL	35 ± 16	30.4 ± 16	69 ± 7*†	44 ± 26‡
STK perfusion (cortex), mL/100 g/min	454.4 ± 20.5	525.0 ± 72.3	283.9 ± 23.8*†	379.3 ± 30.4‡
STK perfusion (medulla), mL/100 g/min	281.7 ± 20.6	512.8 ± 48.4*	130.7 ± 12.4*†	208.6 ± 41.3†
STK glomerular filtration rate, µL/min	170 ± 52	180 ± 63	20 ± 31*†	100 ± 29*†‡
Mean arterial pressure, mmHg	108 ± 12	115 ± 24	171 ± 15*†	160 ± 14*†
IL-1α, pg/mL	9.0 ± 4.6	6.4 ± 0.4	45.9 ± 26.5*	10.2 ± 4.7‡
IL-1β, pg/mL	0.5 ± 0.1	1.9 ± 2.9	4.4 ± 2.9*	0.8 ± 1.3‡
IL-6, pg/mL	6.4 ± 2.2	4.6 ± 2.2	20.5 ± 12.3*†	6.3 ± 1.5‡
IL-10, pg/mL	5.3 ± 0.9	16.9 ± 17.8	5.3 ± 1.5	6.4 ± 2.8
Monocyte chemoattractant protein-1, pg/mL	7.0 ± 1.0	13.1 ± 5.9	28.6 ± 11.2*†	13.8 ± 7.3‡
Tumor necrosis factor-α, pg/mL	8.5 ± 2.9	4.5 ± 2.9	15.9 ± 6.6*†	5.6 ± 1.9‡

n = 8. IL, interleukin; STK, stenotic kidney. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. renal artery stenosis (RAS). One-way ANOVA.

Pericytes Protected the Stenotic Kidney From Apoptosis

Apoptotic activity as assessed by TUNEL and caspase-3 staining indicated almost no apoptotic cells in sham- and sham + pericyte-treated kidneys. RAS markedly increased the number of TUNEL⁺ and caspase-3⁺ cells, which pericyte administration significantly decreased (Fig. 5A).

Figure 5.

A: TUNEL and caspase-3 staining indicated almost no apoptotic cells in sham- and sham + pericyte-treated groups. Renal artery stenosis (RAS) markedly increased the number of TUNEL⁺ and caspase-3⁺ cells, whereas pericytes significantly decreased them. B: minimal spider β-galactosidase (gal)⁺ cells were seen in sham- and sham + pericyte-treated groups, whereas RAS significantly upregulated spider β-gal positivity that was markedly decreased in the RAS + pericyte-treated group. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. RAS. n = 8. t test.

Pericytes Ameliorated RAS-Induced Senescence

Minimal SA-β-gal positivity was seen in sham- and sham + pericyte-treated groups, whereas RAS significantly upregulated cellular senescence and pericyte injection markedly decreased it in the RAS + pericyte-treated group (Fig. 5B).

Pericytes Protected the Renal Microvasculature and Oxygenation

A markedly decreased CD31⁺ staining area (intrarenal microvascular density) was detected in the RAS group, which pericytes significantly increased almost to the level of the sham group (Fig. 6A). Accordingly, the R2* hypoxia index was significantly elevated in the RAS cortex and medulla but distinctly decreased in the RAS + pericyte-treated group to levels comparable to the sham group. Renal expression of HIF-1α was markedly upregulated in the RAS group but was reversed by pericyte delivery. Consequently, gene expression of VEGF and VEGFR was upregulated in the RAS group, and pericytes blunted their expression increment (Fig. 6B). On the other hand, there was no change in VEGF or VEGFR protein expression in the RAS group, yet pericytes increased VEGFR (Flk-1) expression, particularly in the sham + pericyte-treated group (Fig. 6C).

Figure 6.

A: a markedly decreased CD31⁺ area was detected in renal artery stenosis (RAS), which pericytes significantly increased almost to sham levels. Elevated R2* in RAS indicated cortical and medullary hypoxia, which pericytes remarkably decreased. n = 8. One-way ANOVA. B: renal gene expression of hypoxia-inducible factor (HIF)-1α was markedly increased in RAS and improved but not normalized by pericytes. Gene expression of vascular endothelial growth factor (VEGF) and VEGF receptor (VEGFR) was also increased and reversed by pericytes. n = 5. t test. C: pericyte injection greatly increased protein expression of VEGFR (FLK-1), particularly in the sham + pericyte-treated group; VEGF was comparable among the four groups. n = 4. t test. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. RAS. BOLD, blood oxygen level-dependent.

Pericytes Protected the Morphology and Function of the Stenotic Kidney

Body weight was decreased in RAS mice, but pericyte delivery reversed this weight loss (Table 1). RAS also induced hypertension that was significantly attenuated but not abolished by pericytes. Stenotic kidneys had markedly reduced weight and weight ratio to the contralateral kidney, whereas pericytes ameliorated atrophy. Accordingly, stenotic kidney cortical and medullary perfusion was decreased, but only cortical perfusion was restored by pericytes. Interestingly, cortical perfusion was elevated in the sham + pericyte-treated group versus the sham group. Stenotic kidney GFR was reduced and pericytes improved, but not normalized, kidney function (Table 1). However, overall serum creatinine and blood urea nitrogen remained significantly elevated in RAS- and RAS + pericyte-treated groups. Interestingly, blood urea nitrogen was also elevated in the sham + pericyte-treated group.

Pericyte Renoprotective Effects Were Comparable With STCs

Compared with STCs, pericytes had similar effects on ameliorating kidney atrophy, improving cortical perfusion, GFR, and proteinuria. Interestingly, neither pericytes nor STCs affected serum creatinine (Fig. 7).

Figure 7.

Compared with scattered tubular-like cells (STCs), pericytes had similar effects on ameliorating kidney atrophy, improving cortical perfusion, glomerular filtration rate (GFR), and proteinuria. Both pericytes and STCs had no effects on serum creatinine. *P < 0.05 vs. sham; †P < 0.05 vs. sham + pericytes; ‡P < 0.05 vs. renal artery stenosis (RAS). STK/CLK, stenotic kidney/contralateral kidney. n = 8. One-way ANOVA.

DISCUSSION

The present study demonstrates that pericytes possess MSC-like properties and are capable of ameliorating inflammation and oxidative stress in ischemic kidneys, protecting them from tissue hypoxia, vascular loss, fibrosis, and tubular atrophy and eventually preserving stenotic kidney volume and function. Furthermore, the protective effects of pericytes were comparable to those of STCs, a cell type that is known to protect against kidney injury in RAS. These observations suggest that exogenous pericytes may constitute a useful cell-based therapy in chronic kidney disease, particularly ischemic and hypoxic renal injury.

Previous studies have suggested that pericytes exhibit a differentiation potential similar to MSCs (10, 11). In fact, MSCs were proposed to derive from pericytes (10). Although their ultrastructural characteristics have been well studied, pericytes remain relatively poorly defined, with few distinctive markers available for their identification (26). CD146 regulates PDGFR-β activation and is essential for pericyte recruitment (27, 28) but is not widely used in pericyte identification. PDGF-B/PDGFR-β signaling is essential for pericyte proliferation and recruitment to blood vessels (29), and PDGFR-β is one of the best-studied molecular markers expressed in pericytes. NG-2 is expressed on pericyte and oligodendrocyte surfaces during angiogenesis (30), and double PDGFR-β⁺/NG-2⁺ expression patterns are valuable for pericyte identification in tissues (7, 8). α-SMA expression characterizes pericytes and vascular smooth muscle cells but not capillary pericytes (31). Thus, we used PDGFR⁺/NG-2⁺/α-SMA⁻ to discern pericytes from other cells.

Tissue engraftment of exogenously infused MSCs homing to sites of injury and inflammation is the foundation for successful regenerative cell-based therapy (32, 33). In murine and swine RAS models, we have found that MSCs engraft in stenotic kidneys at a higher retention rate than in the contralateral kidney (34), largely due to chemotaxis induced by renal inflammatory cytokines (35, 36). In the present study, we similarly found a higher retention rate of pericytes in stenotic kidneys, which manifested upregulated expression of IL-1α, IL-1β, IL-6, MCP-1, and TNF-α, suggesting that pericyte homing might be regulated similarly to MSCs. Although the majority of injected pericytes were entrapped in the liver and lung, ∼2% were found in the stenotic kidney, which apparently sufficed for their reparative effects. Interestingly, hepatic and pulmonary pericyte retention was also higher in RAS mice than in sham mice, possibly due to cell priming during systemic inflammation (37).

Chronic kidney ischemia may lead to cellular death and kidney atrophy. In the present study, 4 wk after RAS the stenotic kidney shrunk, accompanied by markedly compromised GFR and reduced renal perfusion. In addition, insufficient blood supply decreases oxygen delivery and eventually induces hypoxia (38), as indicated by higher R2* in the stenotic kidney cortex and medulla and upregulated HIF-1α gene expression. Although MSCs had potent antioxidant effects in swine RAS (39), we now show that, similarly, pericytes decrease oxidative stress (DHE staining) and thereby may blunt renal injury and scarring.

The improved microcirculation might have resulted from increased renal VEGF protein expression, possibly bolstered via paracrine mechanisms (37). Interestingly, stenotic kidney VEGF gene but not protein expression was upregulated, possibly due to its proteolytic degradation in chronic ischemia (40). Interestingly, although both MSCs and pericytes preserve the microvasculature, MSCs upregulate VEGF and VEGFR (39), whereas pericytes did not in the present study, implying that their proangiogenic mechanisms might differ. Ischemia and hypoxia downregulate pericyte expression of VEGF, VEGFR, Tie1, and fibroblast growth factor-1 but induce marked pericyte proliferation (41). Pericytes migrate in response to stress or injury to the vascular wall (42), promote endothelial cell proliferation (7), and differentiate into vascular smooth muscle cells, further enhancing vessel maturation and stabilization (42). Thus, their proangiogenic effect might result from a direct regenerative effect to support the vasculature and prevent rarefaction (43). Furthermore, PDGF might be involved in their proangiogenic mechanisms, because a previous study (44) showed that it mediates pericyte recruitment and that blocking PDGFRs inhibits pericyte-induced development of tumor vasculature.

Pericyte delivery reduced both systemic and renal inflammation, as indicated by decreased levels of inflammatory cytokines and macrophage accumulation in stenotic kidneys. Importantly, inflammation participates in the pathophysiology of ischemic kidney injury, and macrophages are important mediators of this process (45). The anti-inflammatory effect of MSCs has been proposed to be mediated through systemic mechanisms rather than direct contact (46). Given their limited retention in the stenotic kidney and the decreased accumulation of macrophages, the anti-inflammatory effect of pericytes might follow a similar pattern to MSCs. Early studies have indicated that pericytes secrete several putative anti-inflammatory factors, including chemokine (C-X3-X motif) ligand 1 and IL-33. IL-33 prevents microglial activation in Alzheimer’s disease (47), and chemokine (C-X3-X motif) ligand 1 promotes an anti-inflammatory microglial phenotype and appears to be neuroprotective (48). In addition, pericytes promote the formation of functionally immunosuppressive regulatory T cell populations and an anti-inflammatory phenotype (49) and decrease immune cell influx to injured tissues (50). Indeed, although ischemia induces the renal release of proinflammatory cytokines such as IL-1, IL-6, and TNF-α (51,52), which, in turn, exacerbate renal injury and fibrosis, pericyte treatment decreased their levels, thereby protecting the stenotic kidney. In vitro, we also found that coincubation with pericytes decreased LDH release from injured cells, confirming their paracrine renoprotective effects. In contrast to MSCs, however, pericytes did not upregulate immune-regulatory and anti-inflammatory IL-10 (53), indicating that they may have fewer immune-regulatory effects.

As we have recently shown (54), RAS also induced cellular senescence in the stenotic kidney, evidenced by increased SA-β-gal staining. Senescent cell accumulation has been implicated in kidney fibrosis through secretion of profibrotic and proinflammatory mediators, so that eliminating them may abrogate the adverse effects of cellular senescence. The ability of pericytes to attenuate cellular senescence might imply this as a potential mechanism for renal protection.

Limitations

Although pericyte treatment improved stenotic kidney oxygenation, single-kidney GFR, inflammation, atrophy, and senescence, it did not attenuate the rise in blood pressure, serum creatinine, and blood urea nitrogen in RAS mice 2 wk after treatment, which might require multiple injections or a higher dose of pericytes. The discrepancy between the improvement in stenotic kidney GFR but not serum creatinine level may be secondary to damage to the contralateral kidney exposed to hypertension. In addition, an observation longer than 2 wk might help uncover their effectiveness in restoring renal function and blood pressure. Future studies are needed to determine the optimal frequency and persistence of the effects of cell delivery over longer periods of time. Although we did not detect cellular rejection, the xenoimmunogenicity of pericytes also warrants additional studies (49). We used as negative controls mice that received the same volume of PBS as that used for pericyte delivery. Ideally, nonrenal cells showing no renoprotection would be used as negative controls, but appropriate cells that would engraft and survive in the kidney are difficult to find. Because our focus was on pericytes, our positive control STC experiments were limited in extent and served as a proof of concept to confirm the reparative properties of pericytes relative to comparable pig-derived xenogeneic reparative cells. Although the body weight of the RAS + pericyte-treated group was similar to that of the sham group, the reason for their greater weight gain compared with sham + pericyte-treated group is unclear and may warrant future studies.

Conclusions

Our study demonstrates a novel cell-based therapy for the injured kidney. The beneficial effect of pericyte delivery appears to be mediated by ameliorating inflammation and oxidative stress in the stenotic kidney and improved tissue hypoxia, vascular loss, fibrosis, and tubular atrophy. Our data may form the basis for pericyte-based therapy, and additional research studies are needed to gain further insight into their role in improving renal function.

GRANTS

This work was partly supported by National Institutes of Health Grants DK120292, HL158691, DK122734, DK122137, and AG062104.

DISCLOSURES

L.O.L. is an advisor to AstraZeneca, CureSpec, Berens Therapeutics, Ribocure Pharmaceuticals, and Butterfly Biosciences. None of the other authors has any conflicts of interest, financial or otherwise, to disclose.

AUTHOR CONTRIBUTIONS

L.O.L. conceived and designed research; T.S. performed experiments; T.S., X-Y.Z., and A.E. analyzed data; X-Y.Z., A.E., Y.J., J.D.K., H.T., and K.L.J. interpreted results of experiments; K.L.J. prepared figures; T.S., X-Y.Z., and A.L. drafted manuscript; L.O.L. edited and revised manuscript; T.S., X-Y.Z., A.E., Y.J., J.D.K., H.T., K.L.J., A.L., and L.O.L. approved final version of manuscript.