Intravenous renal cell transplantation for rats with acute and chronic renal failure

Abstract

Acute kidney injury (AKI) and chronic renal failure (CKD) are the most challenging problems in nephrology. Multiple therapies have been attempted but these interventions have minimal effects on the eventual outcomes, and all too often the result is end-stage renal disease (ESRD). The only effective therapy for ESRD is renal transplantation but only a small fraction of patients receive transplants. In this work we introduce a novel approach to transplantation designed to regenerate kidneys afflicted by severe AKI or CKD: intravenous renal cell transplantation (IRCT) with adult rat primary renal cells reprogrammed to express the SAA gene localized and engrafted in kidneys of rat recipients that had severe AKI or CKD. IRCT significantly resolved renal dysfunction and limited kidney damage, inflammation, and fibrosis. Severe CKD was successfully improved by IRCT using kidney cells from donor rats or by renal cell self-donation in a form of autotransplantation. We propose that IRCT with adult primary renal cells reprogrammed to express the SAA gene can be used to effectively treat AKI and CKD.

acute kidney injury (AKI) complicates chronic kidney disease (CKD), i.e., “acute on chronic,” and it is the most frequent cause of accelerated progression to end-stage renal disease (ESRD) in diabetic nephropathy, the most common cause of CKD (18). Moreover, once ESRD is established, the only specific therapy to get patients off dialysis is kidney transplantation. Unfortunately, ESRD patients face many barriers, in that only 20% are placed on waiting lists for kidney transplantation, and after several years only 10% eventually receive a kidney transplant (35). The rest remain on dialysis and many ultimately succumb to painful complications. AKI is very common (21), constituting an emerging problem in rapidly expanding at risk populations such as the elderly (6), and in patients with diabetes and hypertension (18). Moreover, AKI aggravates the course of CKD, in that accelerated progression to ESRD in CKD is promoted by repeated episodes of AKI, likely from bouts of ischemia and resulting inflammation and fibrosis (18). Accordingly, treatment of AKI is a major goal in nephrology, and over the last 30 yr, multiple therapies for AKI have been introduced, including growth factors, embryonic factors, and mediators of inflammation (5, 14, 17). Regrettably, these interventions have not mitigated the severity of AKI or the progression to ESRD. With the advent of regenerative nephrology, attempts have been made to restore renal function with stem cells, a logical sequel to reports that endogenous stem cells are mobilized to the injured kidney (16, 30), although the latter view has been refuted (8, 23). Multiple stem cell transplantation protocols, including bone marrow-derived mesenchymal stem cells (8), adipose tissue-derived stem cells (9), or amniotic fluid-derived stem cells (12) while showing early attenuation of renal failure, did not result in unequivocal renal engraftment of donor stem cells, or for that matter donor stem cell differentiation into renal cells, which ultimately was the main purpose behind stem cell transplant attempts. We recently demonstrated that the acute phase protein, serum amyloid A (SAA), is critical in tubule formation in cell culture, during tubulogenesis in the embryo and during renal regeneration after an acute injury (19). We now report a novel and successful approach to cell therapy of AKI and CKD based on adult tubular cells reprogrammed with SAA to form tubules in vitro and capable of repopulating denuded and damaged tubules in vivo.

METHODS

Primary renal tubular cells.

Primary renal tubular cells were obtained from normal 175- to 200-g Sprague-Dawley male rats (Harlan, Indianapolis, IN). The rats were killed by removing both kidneys under general anesthesia. Both kidney cortices were minced with scissors in S1 medium (formulation below) and digested with type 4 collagenase (Worthington, Lakewood, NJ), 4 mg/dl, at 37°C in 38% O₂-5% CO₂ for 50 min. The renal tubules were separated by percoll gradient (7), divided into two sets, resuspended in 300 μl of transfection buffer (20 mM HEPES, 142 mM KCl, 6 mM dextrose, 0.7 mM Na₂HPO₄, and 10 μM EGTA), and transfected by electroporation (40 V × 12 ms × 500 ms × 6 pulses; ECM 830 electroporator, BTX, Holliston, MA). Control or SAA1-negative (group “A”) tubules were simultaneously transfected with three vectors: empty vector pcDNA3.1 (30 μg), pAcGFP₁-C₁ [15 μg, green fluorescent protein (GFP) is the cytosolic label used to track cells in vivo; Clontech, Mountain View, CA], and pCruzHA SIRT1 (15 μg, Addgene) (1), which expresses SIRT1, a NAD(+)-dependent deacetylase used to positively influence longevity in transplanted cells (1, 26) in a ratio of 2:1:1. The expressed SIRT1 contains a hemaglutinin (HA) tag useful to track down gene expression (27). For SAA1-positive (group “B”) cells, pcDNA3.1 was replaced with pcDNA3.1-SAA1 plasmid (30 μg), which was manufactured and sequenced in our laboratory, and its construction was previously reported (19). In cell tracking experiments, cells were also transfected with pCAG-TagBFP-V5-NLS [15 μg, this construct induces the exclusive nuclear expression of blue fluorescence protein (BFP) due to its V5 simian virus 5 sequence GKPIPNPLLGLDST and added 3× nuclear localization signal sequences DPKKKRKV] (28, 36).

The cotransfected tubules were cultured in S1 medium: each 2 l contained 10.7 g F-12 HAM, 8.32 g DMEM, 0.29 g l-glutamine, 4.78 g HEPES, 1.7 mg sodium selenite, 0.110 g sodium pyruvate, 3.2 ml phenol red, and pH was adjusted to 7.4 with sodium bicarbonate (Sigma). S1 medium was also supplemented with hepatocyte growth factor (200 ng/ml) and epidermal growth factor (EGF; 400 ng/ml, R&D Systems, Minneapolis, MN). The medium also contained 100 μg/ml hydrocortisone, 35 μg/ml insulin, 32 μg/ml transferrin, 42 ng/ml sodium selenite (Sigma, St. Louis, MO), with 20% FCS, and 75 μg/ml G418 were added after 48 h of culture for selection. The new renal cells growing out of the donor rat tubules initially organized in monolayers, but after 5 days the “B” cells formed new tubules on top of the monolayer (Fig. 1). In preparation for transplantation, male renal tubular cells were lightly trypsinized after 7–8 days in culture, washed in PBS, and injected intravenously in the tail vein of female rats with established AKI or CKD.

Fig. 1.

Reprogrammed renal cells: control rat renal tubular, serum amyloid A (SAA)-negative “A”, cells expand as a monolayer and do not stain with anti-SAA antibody (a). In contrast, SAA-positive (“B”) cells form tubules de novo on top of the monolayer and are SAA positive by immunostaining (b). Nuclear blue fluorescent protein (BFP; e) and cytosolic green fluorescent protein (GFP; g) are strongly expressed in control cells cotransfected with the expression vectors pCAG-TagBFP-V5-NLS and pAcGFP₁-C₁. Nuclear BFP (f) and cytosolic GFP (h) are also strongly expressed in tubule-forming SAA-positive cells (with transfection efficiencies of >70%). Fluorescein uptake by SAA-negative cells is negligible (i). In contrast, tubules formed by SAA-positive cells (j-l) concentrate fluorescein in their lumens. Tubules formed by SAA-positive cells have strong expression of organic anion transporter (OAT)1, the prototypical and highly differentiated OAT (m) (17). SAA mRNA identified by RT-PCR was far more obvious in “B” cells than in “A” cells (n). n, Top: ctrl represents mRNA from “A” cells and “SAAexp” represents mRNA from “B” cells run concurrently with a sample of RNA from “B” cells that was added directly without reverse transcription reaction (−) and an aliquot of amplified SAA plasmid using the same primers (+). Bottom: loading controls using primers for the RAC1 mRNA. SIRT1-HA fusion protein expression in both “A” and “B” cells is demonstrated (o) by probing for the hemaglutinin (HA) tag by immunoblot.

Cotransfected “A” and “B” cells were also cultured on 35-mm glass bottom dishes, and either used for fluorescein uptake studies, or fixed with 4% paraformaldehyde in preparation for immunohistochemistry. Expressed SAA and OAT1 proteins were identified in fixed cells incubated overnight with primary antibodies: rabbit anti-SAA (1:1,000) (19, 20) and rabbit anti-OAT1 (affinity-purified IgG) used at a concentration of 6.5 μg/ml (cat. no. OAT11-A; Alpha Diagnostic International, San Antonio, TX). Cells were rinsed in PBS and labeled with FITC-conjugated goat anti-rabbit IgG (1:200; Jackson ImmunoResearch, West Grove, PA) for 1 h and the nuclear dye Dapi for 20 min. The labeled cells were rinsed with PBS and imaged with an Olympus FV1000 confocal microscope equipped with five lasers (405–990 nm).

For functional studies on tubules formed in vitro, cultured and living cells were first incubated with fluorescein (2 μM, Molecular Probes) added to the culture media overnight. After being rinsed in PBS five times, Hoescht 33342 was added (0.1 mg/ml, Molecular Probes) for 10 min to the media, and cells were again rinsed in PBS and imaged with the confocal microscope as described (19).

Animals.

The 150- to 200-g female Sprague-Dawley rats were anesthetized with intraperitoneal pentobarbital sodium (50 mg/kg) and placed on a homeothermic table to maintain core body temperature at ∼37°C. After ensuring adequate anesthesia, renal ischemia and resulting AKI were induced by occluding both renal pedicles for 50 min with microaneurysm clamps as described (19, 20). There were two groups of rats, 14 in group “A” (received control cells) and 12 in group “B” (received SAA expressing cells). Renal function was monitored by sequential measurements of serum creatinine and blood urea nitrogen with the autoanalyzer of the clinical laboratory at the Indianapolis VA Hospital. For cell infusion studies, 10⁶ cotransfected “A” or “B” cells were lightly trypsinized, washed, suspended in PBS, and infused via tail vein. An aliquot of cells prepared for infusion and passed through the same needle was always recultured to verify viability of the infused cells. The cell viability and growth tests always showed strong cell growth within 24 h of culture. Growth factor and SAA elaboration was determined in media via ELISA according to the suppliers' protocol [Ray Biotech, Norcross, GA for vascular endothelial growth factor (VEGF) and EGF and Immunology Consultant Labs, Newberg, OR for SAA]. For treatment of AKI, IRCT was performed 24 h after ischemia when there was an unequivocal and large increase in serum creatinine and urea nitrogen. In addition, CKD was caused with intraperitoneal cisplatin (1.5 mg/kg every other day × 3 doses) given to two separate groups of seven female rats each. For treatment of CKD, IRCT with “A” (normal cell transfected with control empty vector plasmid) or “B” (normal cells transfected with SAA expressing plasmid) cells was given 3 wk after cisplatin dosing when there was an unequivocal, large, and persistent increase in serum urea nitrogen. A third group (B cispl) with cisplatin-induced CKD was subjected to unilateral nephrectomy 21 days after the last dose of cisplatin. Renal tubules, exposed to cisplatin in vivo, were harvested from each removed kidney, cotransfected with pcDNA3.1-SAA1, pAcGFP₁-C₁, and pCruzHA SIRT1 plasmids, and cultured. The “B” cells, isolated from the B cispl group, were cultured for 8 days in S1 medium in the presence of JAK inhibitor (1 μM; Calbiochem, Rockland, MA); 5 μM CCG1423, inhibitor of Rho A signaling (Cayman, Ann Arbor, MI); 1 μM LY294002, PI3K inhibitor (Wako, Richmond, VA); and 0.1 μM retinoic acid (Sigma) as reported (24) and then were autotransplanted to the same uninephrectomized CKD rat. Two additional control groups of rats receiving three injections of intraperitoneal cisplatin were also included. The experimental time lines were aligned as follows: one group of three rats (cispl) received three injections of platinum and no further treatment during the following 41 days (21 days + 8 days + 12 days) and data were collected during the last 12-day period. The final control group of three cisplatin-treated rats underwent nephrectomy 21 days after the last dose of cisplatin (cispl/neph), received no further treatment during the subsequent 20 days of follow up, and data were collected during the last 12-day period.

Histology and immunohistochemistry.

Kidney sections were fixed in 3.8% paraformaldehyde, paraffin-embedded, and 5-μm sections were obtained for Leder stain to visualize neutrophils and Masson's trichrome to stain connective tissue. In addition, 10-μm sections were obtained for immunohistochemistry. Quantification of histological damage (20, 22) (fraction of tubules with casts, evidence of necrosis and apoptotic cells) was performed on coded sections. Congestion was graded on a scale of 0 (normal) to 4 (congestion observed in all of outer medullary sections). Fixed tissue sections were incubated with primary antibodies: rabbit anti-mouse SAA, rabbit anti-rat OAT1, sheep anti-human Tamm-Horsfall glycoprotein (THP; cat. no. AB733; Millipore, Temecula, CA), rabbit anti-rat thiazide-sensitive cotransporter (TSC; cat. no. TSC11-A; Alpha Diagnostic International), rabbit anti-rat aquaporin-2 (cat. no. AB3274; Millipore), or mouse monoclonal anti-pan keratin (cat. no. 4545; Cell Signaling, Beverly, MA). The secondary fluorescent antibodies were Texas Red-conjugated donkey anti-rabbit or anti-sheep IgG and goat anti-mouse IgG (cat. no. 111-075-045 and 115-075-071, respectively; Jackson ImmunoResearch) and Alexa Fluro647-conjugated donkey anti-sheep IgG (cat. no. A-21448; Invitrogen, Carlsbad, CA). Nuclei were stained with the nuclear dye Dapi (Molecular Probes, Eugene, OR). Images were collected with a Leica DMI 3000B fluorescence microscope and an Olympus FV-1000-MPE confocal microscope and analyzed with MetaMorph software (Universal Imaging, Downingtown, PA).

SAA1 mRNA.

RT-PCR was utilized for amplification of SAA1 mRNA in donor “A” and “B” cells (Fig. 1n). The renal cells were derived from harvested renal tubules and then cultured for 8 days. The cells were mechanically removed from the culture bottles and homogenized in lysis buffer. RNA from donor cells was purified with an RNAqueous-4PCR KIT as recommended by the vendor (cat. no. AM1914; Ambion-Invitrogen). Total RNA was isolated with a purification kit as recommended by vendor (cat. no. 12183-555; Invitrogen) and cleaned with an RNeasy Mini kit as recommended by the vendor (cat. no. 74104; Qiagen, Valencia, CA). For RT-PCR, 2 μg of total RNA were used to synthesize cDNA with an AffinityScript QPCR cDNA Synthesis Kit as recommended by the vendor (cat. no. 600559; Aligent Technologies, Santa Clara, CA). The murine SAA1 mRNA was amplified using the following primers: F1: 5′-CGCCACCATGGAGGGTTTTTTTCATTTGTTCAC, F2: 5′-TACAGGCTAGCGCCACCATGGAGGGTTT, R1/2: 5′-TCAGGTGGATCCCTCAGTATTTGTCAG.

Twenty-five microliters of PCR master solution contained 1× PCR buffer, 0.25 mM dNTP, 0.25 μM primers, 0.25 μl Herculase II DNA polymerase (cat. no. 600675-51; Aligent). cDNA (0.5 μl) was added to each tube before and PCR amplification was conducted: 95°C 5 min 1 cycle, 95°C 20 s; 57.5°C 30 s; 72°C 30 s for 40 cycles, and then at 72°C 7 min for 1 cycle (PCR System 2400, Perkin Elmer, San Jose, CA). The PCR products were separated in a 2.5% agarose gel.

Fluorescent in situ hybridization of the Y chromosome.

Fluorescent in situ hybridization (FISH) was used to localize the Y chromosome in female kidneys 7 days after IRCT with male renal cells as previously reported (34). In short, at termination, recipient kidneys were immediately fixed in 10% neutral formalin, embedded in paraffin, cut into 5-μm sections, and affixed to glass slides. The slides were sequentially placed in xylene for 15 min twice, 100% ethanol for 5 min twice, boiling saline-sodium citrate (SSC) buffer for 10 min, cooled down to room temperature, and washed with distilled water. The sections were then digested with 0.4% pepsin, 0.9% NaCl, pH 1.5, at 37°C for 50 min, washed with distilled water, immersed in 2× SSC for 10 min twice, and air dried. FISH was conducted with the fluorescent-labeled rat Y chromosome probe (Rat Idetet Chr Y Paint probe red, ID 556; cat. no. IDRR1070–0111 Ex:548; Em:573, from ID Labs Biotechnology, London, ON, Canada) diluted 1:10, and 5 μl were applied to the prepared kidney section. The slide was covered, and kidney DNA and probe were denatured at 69°C for 2 min, and then hybridized overnight at 40°C. The slide was then uncovered and placed in warmed 0.4× SSC with 0.3% NP-40 (70°C for 2 min). The slide was then incubated in 2× SSC solution at room temperature for 1 min. The section was then washed with distilled water, air dried, mounted with DAPI (cat. no. F/203/F204; Insitus Biotechnologies, Albuquerque, NM), covered, and imaged on a fluorescent microscope.

Immuno (Western) blotting.

Cells were homogenized in 25 mM Tris, pH 7.6, 150 mM NaCl, 1% deoxycholate, 1% P-40, 0.1% SDS, and 2× halt protease inhibitor cocktail (Thermo Scientific, Rockford, IL) and adjusted to a protein concentration of 2 mg/ml. The homogenates (20 μg) were fractionated by electrophoresis through 16.5% polyacrylamide Tris-Tricine gels. After electrophoresis, proteins were transferred to a nitrocellulose filter. Blocking was carried out in 1% casein, 1× PBS for 1 h. Incubation with primary antibodies diluted in 1× PBS was for 1 h; primary antibodies were rabbit anti-influenzae HA (1:250; Santa Cruz Biotechnology, Santa Cruz, CA) and mouse anti-actin (1:1,000; clone AC-40, Sigma). The filter was then washed in 1× PBS and incubated with secondary antibodies diluted in 1× PBS for 1 h. Secondary antibodies were IRDye 680 goat anti-rabbit IgG (1:15,000; Li-Cor Biosciences, Lincoln, NB) and IRDye 800 CW goat anti-mouse IgG (1:20,000; Li-Cor Biosciences). After being washed in 1× PBS, the filter was scanned using an Odyssey Infrared Imaging System (Li-Cor Biosciences) for visualization of immunoreactive proteins.

Statistics.

All image quantification was performed on coded images. Data are expressed as means ± 1 SE. ANOVA was used to determine whether differences among mean values reached statistical significance. Tukey's test was used to correct for multiple comparisons. The null hypothesis was rejected at P < 0.05.

Animal use statement.

The experiments were conducted in conformity with the “Guiding Principles for Research Involving Animals and Human Beings.” The investigations were approved by the Institutional Animal Care and Use Committee of Indiana University School of Medicine.

RESULTS

Creation of SAA reprogrammed primary tubular cells.

We followed our finding that SAA promoted tubule formation in cultured cells, during embryogenesis, and recovery from AKI (19), by testing SAA in renal injury. Kidney tubules were harvested from normal male rats, transfected by electroporation, and divided into two groups, “A” (control) and “B” (SAA expressing; see methods). “A” tubules received pcDNA31.1, (empty vector), pacGFP1-C1 (GFP is the cytosolic label), pCruzHA SIRT1 expresses SIRT1, an NAD(+)-dependent deacetylase (26, 27), and in specific cell tracking experiments pCAG-TagBFP-V5-NLS (targets BFP to cell nuclei) (36). “B” tubules received same plasmids and total amount of DNA, except that pcDNA31.1 was replaced by pcDNA31.1.SAA1 (expresses SAA1 protein) (19). The tubules were cultured in S1 medium and new cells grew out from the original tubules within 48 h and expanded in monolayers from which tubules emerged de novo after 72 h on culture. Tubule formation in SAA-positive “B” cells was robust and fourfold higher than in SAA-negative “A” cells. These findings are similar to our prior results with reprogrammed NRK42E cells (19, 20) (Fig. 1). The branching tubules constituted of SAA-positive (“B”) cells expressed SAA protein and RNA (Fig. 1, b and n), whereas control “A” cells were SAA-negative. Both “A” and “B” cells expressed cytosolic GFP, nuclear BFP, and SIRT1 (Fig. 1, c-h and o). These new tubules were differentiated and functional, as demonstrated by uptake of the weak acid fluorescein (Fig. 1, i-l). Tubular differentiation was further verified by expression of the transporter OAT1, a sensitive differentiation marker in renal tubules (19) (Fig. 1m). Cell secretion of VEGF, EGF, and SAA into media was detectable in all cases with no difference in VEGF and EGF secretions between “A” and “B” cells (Table 1).

Table 1.

Secretion into the media of VEGF, EGF, and murine SAA by A and B primary renal cells

Factor	A Cells	B Cells	P Value
VEGF, pg•ml−1•μg protein−1	37.74 ± 2.15	34.18 ± 1.07	0.16
EGF, pg•ml−1•μg protein−1	0.20 ± 0.004	0.18 ± 0.009	0.10
SAA, ng/ml	0.10 ± 0.03	12.97 ± 0.71	0.02

Values are means ± SE. VEGF, vascular endothelial growth factor; EGF, epidermal growth factor; SAA, murine serum amyloid A1 protein.

Renal cell transplant in renal ischemia-reperfusion injury.

Two groups of female rats subjected to ischemia-reperfusion (I/R) injury for 50 min at 37°C sustained severe AKI as indicated by the rapidly rising serum creatinine and blood urea nitrogen levels (BUN; Fig. 2). SAA-negative (“A”) and -positive (“B”) renal tubular cells, 10⁶ each, were then given intravenously in the tail vein of rats from the two groups. The cells were infused 24 h after I/R when acute renal failure was already well-established. Infusion of SAA-negative “A” cells did not appreciably change the severity of their renal failure, and their uremic picture remained unabated, with their serum creatinine and BUN levels increasing further over the subsequent 24-h period (Fig. 2). The abrupt surges of serum creatinine and BUN in rats receiving “A” cells were indistinguishable from those seen in rats subjected to the same degree of I/R without cell infusion (4 ± 0.2 mg/dl 48 h postischemia). In marked contrast, renal failure in the group of rats given SAA-positive “B” cells was less severe, improved rapidly, and their recovery was sustained during the period of observation. The dramatic recovery induced by “B” cells is further exemplified by the fact that all 12 rats infused with “B” cells survived AKI, but of the14 rats receiving “A” cells, 5 died within days as a result of AKI, P = 0.02. In addition, histological evidence of injury 48 h postischemia was markedly attenuated in the “B” cell group (Fig. 2, d-e).

Fig. 2.

Effects of intravenous renal cell transplantation (IRCT) on renal function and structure: two groups of female rats sustained ischemia-reperfusion injury (IRI) at time 0 and then received IRCT with control “A” (n = 12) or SAA-expressing “B” renal cells (n = 12). Serum urea nitrogen and creatinine reported the severity of the renal failure 1 day after cell injection (a and b). IRCT with SAA-positive cells caused a substantial fall in both urea and creatinine by the following day, whereas IRCT with control cells had no effect in higher levels of these 2 indicators. Mortality was significantly lower in rats that received “B” cells (compared with “A” cells; c). Renal structure was visualized with periodic acid Schiff (PAS) stain and revealed that rats post-IRI treated with “A” cells sustained severe tubular injury with dilatation, necrosis, cast formation, and the presence of free cells in tubular lumens (d). In marked contrast, tubule structure of kidneys from the group of rats that sustained the same degree of IRI but were transplanted with SAA-positive “B” cells was much better preserved with minimal tubular damage and a few casts (e). Quantification of multiple images demonstrated significantly less necrosis, cast formation, and apoptosis in kidneys from “B” cell-treated rats (f).

Not only infused “B” cells improved renal function, but those rats recipient of “B” cells maintained a remarkable preservation of their renal structure (Fig. 3). Thus, 48 h after I/R, and 24 h after infusion of “A” cells, there was a very conspicuous broad band of vascular congestion along the entire ischemic renal outer medulla (Fig. 3a). Furthermore, detailed views of the outer medulla at low microscope power reveled red cells distinctly stacked up in medullary vessels along with heavy neutrophil infiltrates. On the other hand, the renal macroscopic and microscopic appearance of rat kidneys injected with “B” cells was nearly normal after 24 h postinfusion, in that only very limited vascular congestion and neutrophils were found by comparison (Fig. 3b). Leder stain verified the massive neutrophil renal influx in rats given “A” cells, while kidneys from rats treated with “B” cells had only occasional neutrophils. Mean leukocytes/hpf was 12.3 ± 2.5 in the “A” group vs. only 1.1 ± .3 in the “B” group (P = 0.006). Moreover, 3 wk after cell infusion, renal peritubular, or interstitial, fibrosis was clearly discernible after cell transplantation with “A” cells, whereas minimal of fibrosis was found in kidneys treated with “B” cells (Fig. 3, f-i). Quantification of renal fibrosis (mean blue pixel density in 14 sections) showed that in the “B” cell group (93 ± 14), fibrosis was significantly less (P = 0.039) than in the kidneys of rats treated with “A” cells (137.9 ± 13).

Fig. 3.

Renal pathology in female rats with acute kidney injury (AKI). Rats sustained AKI from ischemia-reperfusion (I/R) for 50 min and received IRCT 24 h later with either control SAA-negative (“A”) cells (a, d) or with SAA-positive (“B”) cells (b, e). Representative images of sectioned kidneys show severe vascular congestion in the outer medulla of kidneys that received control “A” cells (a). Inset: microvasculature is congested with erythrocytes and leukocytes, more evident at higher magnification. In contrast, vascular congestion was barely detectable in the kidneys that received SAA-positive “B” cells (b). The bar diagram (c) summarizes the degree of vascular congestion, represented by an arbitrary scale from 0 to 4, and it shows far more severe congestion in rats that received control A cells (n = 40 for both). Leukocytes, visualized by Leder stain, were significantly more abundant after IRCT with “A” (d) than with “B” (e) cells. D, inset: aggregated leukocytes at the arrow. Fibrosis (blue stain) was also significantly greater 3 wk postischemia in the “A” group (f, h) than the “B” group (g, i) both in renal cortex (f, g) and renal medulla (h, i).

Cell engraftment.

The current view is that after AKI, stem cell transplants improve renal function acutely by paracrine action, a conclusion largely based on the relatively poor engraftment characteristic of stem cell transplantation (3, 4, 10). However, in the long term, successful cell engraftment may yield superior structural and functional results. Consequently, cell engraftment 7 days posttransplant was evaluated by independent and complementary approaches. The cultured donor renal cells were of epithelial origin as indicated by uniform immune labeling with pan-keratin antibody (not shown). In Fig. 4 is shown that at the time of transplant, donor renal cells were positive for organic anion transporter 1 (OAT1; proximal tubule), THP (thick ascending limb), and aquaporin-2 (collecting tubule) markers, while there was a paucity of thiazide-sensitive cotransporter-positive cells (distal convoluted tubule; Fig. 4). In the kidney, 7 days after IRCT, transplanted GFP-positive tubular cells colocalized with OAT1 and THP, indicating preferential cell engraftment in proximal tubule and thick ascending limb, respectively (Fig. 4). Donor GFP-positive cells obtained 1 and 7 days post-IRCT are shown in recipient kidneys (Fig. 5, a-d). Quantification of GFP-positive cells engrafted in renal tubules (per ×60 microscopic field) showed initial equal numbers in kidneys from “A” (control) and “B” (SAA expressing) groups. However, by 7 days “A” cells had decreased while “B” cells increased, P = 0.001 (n = 11 for each), and at 21 days, the difference between “A” and “B” groups was even greater (P = 0.0000007; n = 3). In contrast, GFP-positive donor cells were only rarely seen in lungs, spleen, and liver in both groups. We also employed FISH to identify the Y chromosome of transplanted male donor cells in female kidneys 7 days post-IRCT (Fig. 5). FISH revealed the presence of Y chromosomes in male kidney tubules (Fig. 5e) as well as in the tubules of female kidney recipients of male tubular cells (Fig. 5, g and h) but not in normal female kidneys (Fig. 5f). In short, independent protocols indicate that donor renal cell engraftment followed the IRCT procedure.

Fig. 4.

Characterization of primary cells: immunostaining of cells after 7 days in culture (left vertical panels) demonstrated that formed tubules in vitro expressed OAT1, Tamm-Horsfall protein (THP), and aquaporin 2 (AQP), but very few cells expressed the thiazide-sensitive cotransporter (TSC). The fraction of cultured GFP-positive, SAA-expressing “B” cells that colocalize with specific tubular markers is shown in the inset, indicating that the largest fraction of transfected cells was either OAT- or THP-positive. In renal tubules, 7 days after cell transplantation, many cells coexpressed both OAT1 (red) and GFP, or THP (red) and GFP while coexpression of TSC or AQP with GFP was not observed. Thus, IRCT was performed with a mixture of tubular cells, but engraftment was primarily observed in the proximal tubule and thick ascending limb, known areas of cell death after ischemia (22, 33). Insets of tubule cells at dotted arrows demonstrate expression of OAT1 or THP (red) in the same cells as GFP.

Fig. 5.

Renal cell engraftment of tubular “B” (SAA-expressing) cells 7 days post-IRCT. The originating primary tubular cells were transfected with plasmids designed to express SAA and cytosolic GFP. Twenty-four hours after infusion, GFP-positive cells are seen in the interstitium and tubules in “A” (control) and “B” groups (a, b). Seven days after IRCT, multiple labeled cells were seen lining recipient renal tubules after IRCT with “B” (d) but fewer “A” cells were detected in this section (c). Renal cell engraftment was also verified by fluorescent in situ hybridization (FISH) 7 days post-IRCT. FISH was conducted with a red fluorochrome-labeled DNA probe specific for the rat Y chromosome. Four representative conditions are depicted: normal male kidney positive control (e), normal female kidney negative control (f), AKI female kidney transplanted with control SAA-negative “A” cells (g), and AKI female kidney transplanted with SAA-positive “B” cells (h). The Y chromosome-positive donor nuclei (red spots) were found integrated and adjacent to the host female cell nuclei. Male (FISH-positive) and female (FISH-negative) nuclei were distributed along renal tubular outlines. Quantification of engrafted GFP-positive “A” and “B” cells (in a total of 141 images) is shown in i.

Renal cell transplant in CKD.

Renal cell transplantation and successful engraftment have the potential to improve CKD, a condition characterized by progressive loss of functional renal tissue. Therefore, we tested the role of IRCT in a model of CKD caused by cisplatin toxicity. Three groups of rats were administered intraperitoneal cisplatin, 1.5 mg·kg⁻¹·day⁻¹, every other day for three doses; 3 wk later the rats had acquired severe renal failure and their mean BUN was 116 ± 7 mg/dl. The rats were then divided into three groups, and two groups received IRCT. Rats in the “A” group received SAA-negative “A” cells and rats in the “B” group received SAA-expressing “B” cells, and their renal function was monitored for 12 more days posttransplantation (Fig. 6a). IRCT with “A” cells did not change the established uremic state of “A” rats. However, notwithstanding a slower response to IRCT in CKD than in AKI, significant resolution of the uremic picture was detected 6 days posttransplantation in rats of the “B” group. More importantly, the improvement in CKD was marked and sustained during the entire 12-day period of observation post-IRCT (Fig. 6a). The third group of rats was not given any cells at all and their severe uremic state was comparable to the rats given “A” cells. In the same Fig. 6a is shown that renal autotransplantation with SSA-expressing renal cells also improved very severe CKD in cisplatin-intoxicated rats that were then subjected to unilateral nephrectomy. There were two additional groups of rats given intraperitoneal injections of cisplatin, 1.5 mg·kg⁻¹·day⁻¹, every other day for three doses. One group of rats had a unilateral nephrectomy 21 days after cisplatin injections, their tubules were harvested, transfected with SAA-expressing plasmid, and cultured for 8 days. The resulting SAA-expressing renal cells were then isolated and given back intravenously to the donors (autotransplantation, B cispl). Following IRCT with their own cells, the severe renal failure, BUN averaged 156 ± 8 mg/dl, rapidly improved over the next 12 days post-IRCT. The other group of rats was also given three intraperitoneal injections of cisplatin, and 21 days later was subjected to unilateral nephrectomy. The rats were then followed for 8 more days to mimic the required period of in vitro cell growth, but were not given IRCT. At this time point, renal failure was very severe, BUN averaging 138 ± 6 mg/dl, which continued to gradually increase over the next 12 days of observation (Fig. 6A). All rats were killed at the end of the 12-day observation and transplanted cell engraftment in renal tubules was confirmed by confocal fluorescence microscopy of donor cells expressing cytosolic GFP and nuclear BFP in a representative “B” cells recipient kidney (Fig. 6b). The purpose of having simultaneous nuclear and cytosolic fluorescent protein expressions was to strengthen donor cell identification, in that concurrent visualization of properly localized nuclear BFP and cytosolic GFP is unlikely to occur by mere transfer of fluorescence markers from donor to host cells. In addition, donor SAA-positive “B” cells were easily identified by fluorescence microscopy in all rats (Fig. 6, c-h). SAA is shown in red in two representative rats (Fig. 6, c and f), their cellular GFP in Fig. 6, d and g, and their Dapi-stained nuclei in Fig. 6, e and h. The renal histology showed that kidneys from rats in the “A” group had advanced tubular atrophy and cast formation (Fig. 6i; no different from that without cell treatment, data not shown), whereas kidneys from rats in the “B” group showed remarkable preservation of the tubular architecture (Fig. 6j). Masson's trichrome stain revealed extensive peritubular and glomerular fibrosis in CKD rats infused with “A” cells, as shown in a representative section (Fig. 6l), and remarkable ablation of glomerular and peritubular fibrosis in CKD rats infused with “B” cells (Fig. 6m). IRCT with “A” or with “B” cells also had substantial differential effects on body and kidney weights of recipient rats: the initial body weights of rats receiving “A”, “B”, and cisplatin “B” cells were similar, 156 ± 7, 147 ± 1, and 152 ± 7, respectively (g; P = NS). In contrast, body weights 12 days after cell transplant were 185 ± 3, 202 ± 8, and 200 ± 3, respectively (significantly higher for both B cell groups vs. control “A” cells, P = 0.021). Furthermore, terminal kidneys weights for “A” and “B” rats were also different, 0.78 ± 0.01, 0.72 ± 0.01, and 0.74 ± 0.01, respectively, for the 3 groups (significantly lower for both “B” cell groups vs. control “A” cells, P < 0.05).

Fig. 6.

Effects of IRCT in chronic kidney disease (CKD). Three groups of rats with CKD following 3 intraperitoneal injections of cisplatin given on alternate days were studied. Twenty-one days after cisplatin injections, 2 groups of rats received IRCT with either control SAA-negative (control, blue line during 12 days posttransplant) “A” or SAA-positive “B” cells from (brown line during 12 days posttransplant) normal rats and then followed 12 more days. The third group was monitored for 33 days after cisplatin injections and received no cells at all (gray line, during last 12 days of observation synchronized with the other 2 groups). Two additional rat groups were given 3 intraperitoneal injections of cisplatin given on alternate days and were uninephrectomized 21 days later. Tubules isolated from kidneys in one group of rats were transfected with SAA plasmid and cultured for 8 days. The cultured SAA-expressing “B” cells were then given back intravenously to each respective donor rat (black line, during the last 12 days following IRCT). The last group was uninephrectomized 21 days after the last cisplatin injection, monitored for 8 more days, and data were collected during the last 12 days of observation (green line, during the last 12 days of observation). Engraftment of renal cells expressing cytoplasmic GFP and nuclear BFP is shown in 12 days post-IRCT (b). Immunoreactive SAA (red) is also evident in tubules of kidneys treated with “B” cells (c and f). These cells are also GFP-positive (d and g). All nuclei are stained with Dapi and shown in e and h. Representative microscopic images of kidneys 33 days after cisplatin injections and 12 days post-IRCT: PAS stain revealed marked tubular dilatation and intratubular casts and cells in the kidney of a rat infused with “A” cells (i), whereas the kidney infused with SAA-positive “B” cells from normal (j) or CKD (k) rats had significantly less histological evidence of renal injury. Kidney sections stained with Masson's trichrome showed extensive glomerular and peritubular fibrosis (blue) of the kidney infused with control “A” cells (l), and much less peritubular fibrosis, and nearly normal glomeruli, in the kidney of the CKD rat infused with SAA-positive “B” cells from normal (m) or CKD (n) kidneys.

DISCUSSION

AKI, either by itself, or when complicating and aggravating CKD is one of the most challenging clinical problems in nephrology. The evolution of established AKI is generally unpredictable and its course ranges widely, from minimal effects to causing complete renal failure or even ESRD (18, 21, 31). There is no effective treatment for AKI (21). CKD is even more problematic, in that only very modest gains have been achieved even when targeting known causes, and many patients wind up with ESRD and on dialysis (13). Once on dialysis, renal transplantation remains the only effective means of freeing ESRD patients from dialysis dependency. Unfortunately, the demand for kidneys vastly exceeds supply, so kidney transplantation is only available to a small minority of dialysis patients (35). Consequently, numbers of CKD and ESRD patients continue to climb and so does the enormous economic cost required for their care (32). Accordingly, new tactics are badly needed to treat AKI and to limit CKD. In this report, we describe the role of IRCT in rats with AKI and CKD and renal cell autotransplantation in rats with CKD. We successfully performed intravenous renal cell transplantation (IRCT) with rat primary adult renal tubular cells that were reprogrammed to express the murine isoform of SAA. New renal cells grew out of harvested renal tubules when cultured for 7–8 days, and the SAA-positive cells formed more new tubules in vitro; the basic element of redifferentiation. SAA was chosen to reprogram and redifferentiate cultured renal cells because of its powerful redifferentiation and tubulogenic properties (19). Indeed, current and past results (19, 20) strongly indicate that SAA expression in donor cells is a critical element for tubular formation and renal function recovery. For instance, IRCT with rat SAA-expressing NRK52E cells, an otherwise poorly differentiated cell line, drastically improved renal function in multiple models of AKI, whereas IRCT with wild-type NRK52E cells had no effect. These differential effects on AKI between SAA-positive and SAA-negative NRK52E cells were similar to the current report with primary renal cells (20). The critical distinction between the two IRCT studies is the use of normal adult renal tubular cells and their remarkable effectiveness in advanced AKI and also in CKD, which makes IRCT amenable to clinical applications in AKI and CKD. Although atherogenic or carcinogenic effects of SAA were not evaluated in the present study, the very low levels of SAA secretion by “B” cells (Table 1) are unlikely to result in systemic effects. The renal tubular cells used for IRCT were derived from cultured renal tubules harvested from normal rats or from rats with CKD and reprogrammed with the SAA gene. The cell yield after 8 days in culture was sufficient to transplant six rats from a single normal donor rat. IRCT with “B” cells improved renal function within 24 h, a relatively early action consistent with paracrine or endocrine effects (2, 31), perhaps from one of the many potential factors released by cultured cells (29). In any case, donor renal tubular cells reached the kidney via the circulation, became engrafted, and proliferated in the host damaged tubules. Once they had reached the kidneys, donor tubular cells multiplied and filled empty spaces presumably left by damaged or dead host renal tubular cells. The donor cells were an admixture of tubular cells expressing OAT1, THP, and aquaporin-2, while TSC-expressing cells were poorly represented. The engrafted donor cells continued to express cell markers of the proximal tubule and thick ascending limb, suggesting localization to damaged proximal tubule and thick ascending limb, segments severely affected in I/R injury (22, 33). It is not clear whether poor distal nephron engraftment occurred because of a low number of specific donor cells, or lower cell deletion in these two segments, which limited space for host transplanted cells.

IRCT with reprogrammed but otherwise normal primary renal cells is a feasible method for clinical use, and its foundation and applicability are based on solid grounds. Following a long period of conflicting investigations, the prevailing view is that proliferation and expansion of residual resident adult renal epithelial cells are the main mechanisms of renal repair in AKI (8, 15, 23, 33). In short, endogenous stem cells do not seem to play a role in renal regeneration (10). Nevertheless, stem cell transplantation has been attempted to promote renal regeneration and was found to improve renal function in AKI (10). However, the benefit seemed to be derived from paracrine or endocrine activity and not from cell engraftment, as stem cells apparently do not localize to damaged tubules (3). Hence, it is hard to estimate the benefit of stem cell transplantation after the paracrine effects fade away, in that even when the improvement is reportedly sustained, the observation time is usually short in AKI, and within the time frame of substantial spontaneous organ recovery (20). Furthermore, to our knowledge there is no evidence that stem cell transplantation improves CKD as documented for IRCT with “B” cells. We tested IRCT on a severe model of CKD with advanced waste retention and renal structural damage. Moreover, we were able to perform renal cell autotransplantation that also improved CKD. These “B” (SAA-positive) cells were grown in sufficient numbers from a nephrectomized kidney to affect significant correction of the uremic state. We found that IRCT improved uremia, restored tubular and surprisingly glomerular integrity, and limited renal fibrosis. These changes were accompanied by striking restoration of growth rates in CKD rats, in that rats that received IRCT with “B” cells grew by 37% from the time of injury compared with only 18% growth in rats that received “A” cells. Therefore, we propose that AKI and CKD treatment with IRCT based on reprogrammed primary renal cells is a very effective therapy and compares quite favorably with stem cell transplantation. Moreover, IRCT with primary renal cells is not plagued by a myriad of ethical and legal issues, including property rights (25), and it also has the potential, in selected cases, to be used as a form of renal saving autotransplantation.

GRANTS

This work was supported in part with funds from the National Institutes of Health (5R01DK082739) and Paul Teschan Research Fund of Dialysis Clinica to K. J. Kelly and the VA Merit Review program to J. H. Dominguez.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: K.J.K., M.W., S.Z., and J.H.D. conception and design of research; K.J.K., J.Z., M.W., S.Z., and J.H.D. performed experiments; K.J.K., J.Z., M.W., S.Z., and J.H.D. analyzed data; K.J.K., S.Z., and J.H.D. interpreted results of experiments; K.J.K., J.Z., and S.Z. prepared figures; K.J.K. and J.H.D. drafted manuscript; K.J.K. and J.H.D. edited and revised manuscript; K.J.K. and J.H.D. approved final version of manuscript.