Restoration of tubular epithelial
cells during repair of the postischemic kidney occurs independently of bone marrow-derived stem cells (original) (raw)
Ischemic injury results in severe tubular damage followed by repair. Kidney ischemia/reperfusion (I/R) results in disruption of the actin cytoskeleton in damaged tubular cells, which is reversible if the cell survives the insult (3). The actin cytoskeleton was identified by staining with phalloidin conjugated to the cyanine dye Cy3 (phalloidin-Cy3). Two days following unilateral ischemia, there was loss of the brush border of proximal tubular cells of the S3 segment in the postischemic kidney but not in the contralateral control kidney (Figure 1A). Extensive loss of epithelial cells could be seen in some tubules after ischemia. Seven days following injury, many of the tubules had recovered an apical actin cytoskeletal staining pattern, indicative of restoration of the brush border, and 3 weeks following I/R injury, the lost tubular cells had been almost completely replaced by new cells (Figure 1A).
Characterization of I/R injury in chimeric mice. (A) Phalloidin staining of actin cytoskeleton in sham-operated (control) kidneys and at 2, 7, and 21 days following I/R injury. Note the loss of the tubular cell brush border at 2 days and the denuded tubules (arrow) and the progressive restoration with time of both apical actin staining and tubule integrity. (B) PAS-stained sections at 2 and 7 days following I/R injury. Although the necrotic intratubular debris (arrows) is widespread at 2 days, the tubular architecture is significantly restored at 7 days. (C) PCNA staining of tubules in sham-operated (control) and ipsilateral kidney 2 days following I/R injury. (D) Outer medulla intratubular proliferation as assessed by counting mitotic cells in PAS-stained sections and by anti-PCNA immunofluorescence at 0, 2, and 7 days following I/R injury to the kidney. (E) Plasma creatinine levels at 0, 2, and 7 days following bilateral I/R renal injury. Scale bars: 50 μm.
I/R injury resulted in characteristic necrotic and apoptotic changes in the tubules of the outer medulla, followed by intense proliferation and recovery of tubular integrity (Figure 1B) (3, 18). In addition to identifying mitotic nuclei, we stained sections for nuclei expressing the protein proliferating cell nuclear antigen (PCNA), in order to label and quantify proliferating cells (Figure 1C). Tubular cell proliferation was greatly increased (63.7% ± 4.7% PCNA-positive nuclei) in the outer medulla of the recovering kidney compared with the sham-operated control kidney (0 hours) (Figure 1D). Tubular cell proliferation also remained elevated (8.1% ± 1.3%) at 7 days. Bilateral ischemic injury resulted in a rise in plasma creatinine levels from baseline of 0.1 mg/dl to a peak of 1.7 mg/dl at 48 hours (n = 4 per time point), followed by complete recovery to baseline at 7 days (Figure 1E).
Enhanced GFP is not detected in tubular cells of enhanced GFP chimeric mice following I/R injury. We generated chimeric mice with donor bone marrow expressing enhanced GFP (EGFP). In the donor, the EGFP gene was driven by the chicken β-actin promoter and was expressed in all tubular cells (19). I/R injury was induced in male mice 6 weeks following bone marrow transplantation. Bone marrow cells and buffy coat leukocytes in the recipients were analyzed by flow cytometry and compared with cells from the donor mice (Figure 2A). The proportion of bone marrow cells of recipients with increased fluorescence from EGFP (57.6% ± 3.1%; n = 12) was comparable to that of donor mice bone marrow cell expression (59.6% ± 4.2%; n = 3). The proportion of total blood leukocytes with EGFP fluorescence was somewhat higher in both chimeras and donor mice (73.5% ± 3.5%, n = 12; and 77.4% ± 4.7%, n = 3, respectively). The fact that the number of fluorescent cells in bone marrow was not an even higher percentage of total cells likely reflects the previously reported lack of expression of EGFP in cells of the erythroid lineage (19). Myeloid leukocytes (77.7% ± 3.3%) and circulating lymphocytes (74.0% ± 2.5%) in the chimeras also expressed EGFP, which confirmed that chimerism was not restricted to 1 leukocyte lineage (Figure 2B). In addition, when tissue sections of spleen and thymus from the chimeric mice were analyzed, nearly all cells expressed EGFP (Figure 2C). These data indicate that multipotent hematopoietic stem cell progenitors were transplanted and engrafted into the bone marrow.
Confirmation of chimerism in mice transplanted with bone marrow from EGFP-chimeric mice. (A) Top panels: Representative FACS analysis for EGFP fluorescence of bone marrow from wild-type mice (left) and a chimeric mouse at 6 weeks after bone marrow transfer (right). Bottom panels: FACS analysis for EGFP fluorescence of blood leukocytes from wild-type mice (left) and chimeric mice at 6 weeks (right). (B) Representative analysis of leukocytes from EGFP chimeras for EGFP fluorescence and expression of the myeloid lineage marker CD11b (left) or lymphoid lineage markers B220, CD4, and CD8 (right). Note that in each case, the majority of circulating leukocyte populations express EGFP at high levels. (C) Tissue sections of spleen and thymus in chimeric mice showing widespread EGFP fluorescence. Scale bar: 50 μm.
In an attempt to detect cells within the tubule that expressed EGFP in these chimeric mice, we examined sections from the ipsilateral and contralateral kidneys of mice at 2 days (n = 4) and 7 days (n = 4) after ischemia at high power (magnification, ×400). We found no examples of tubular cells expressing EGFP at these time points (Figure 3). By contrast, analysis of kidney sections from donor EGFP mice confirmed that tubular cells strongly expressed EGFP (Figure 3C), and, therefore, tubular cells in the chimeric mice would be expected to be clearly visible if they were derived from BMSCs from the donor mouse.
Tubular cells in EGFP chimeras do not express EGFP following I/R injury. Photomicrographs of corticomedullary regions of the contralateral kidney (A) and kidney 7 days after I/R in chimeric mice (B). Note bright EGFP in nontubular cells but only dull autofluorescence in tubular cells. (C) Photomicrograph of kidney from an EGFP donor mouse as positive control showing bright EGFP in tubular cells. Scale bar: 50 μm.
Following I/R injury, EGFP is detected in peritubular cells coexpressing leukocyte markers and a small population of cells lacking CD45 and expressing endothelial markers. The postischemic kidneys of EGFP chimeras had an increased number of fluorescent cells (Figure 3B). In order to determine the phenotype of these cells, we immunolabeled tissue sections for leukocyte markers (B220, CD11b, and CD45) and markers of endothelial cells (vWF and CD31), then examined them by confocal microscopy. Not surprisingly, most EGFP-positive cells stained positively for leukocyte markers. Nevertheless, small numbers of EGFP-positive cells (0.5% ± 0.6%) did not express the leukocyte common antigen CD45 (Figure 4A) 7 days following I/R injury. In areas, particularly in the outer medulla, where extensive tubule repair had occurred by 7 days following I/R injury, some EGFP cells also expressed CD31, a marker of endothelial cells (Figure 4B). We examined sections by confocal microscopy to confirm colocalization (Figure 4B). In sections examined by confocal microscopy, vWF, another marker of endothelial cells, was also found to be coexpressed in a small subset of EGFP-positive cells (Figure 4C). EGFP-positive endothelial cells were found only in peritubular capillaries. Neither capillary loops of glomeruli nor arterioles showed any evidence for EGFP positivity. Importantly, in the contralateral kidneys, very few endothelial cells expressing EGFP could be seen (data not shown). In order to determine the proportion of endothelial cells expressing EGFP, serial high-power fields (HPFs) in the cortex were scored for green fluorescent cells that also expressed either vWF or CD31 (Figure 4, B and C). As quantitated in Figure 4D, 0.3% ± 0.3% of endothelial cells in contralateral kidneys and 1.6% ± 0.4% in 7-day postischemic kidneys coexpressed vWF and EGFP. Similar observations were made for CD31-positive cells (Figure 4D). Together, these data suggest that endothelial cell replacement by BMSCs or fusion with bone marrow–derived cells occurs at a low level in response to the vascular injury resulting from ischemia. Sections were also labeled with the fibroblast marker α-SMA. Despite an expanded population of α-SMA–positive cells, we found no evidence by confocal microscopy for EGFP-expressing fibroblasts in postischemic kidneys (data not shown).
Bone marrow EGFP-expressing cells lack leukocyte markers and acquire characteristics of peritubular endothelial cells 7 days following I/R injury. (A) Fluorescence images of 7-day postischemic kidney showing interstitial cells expressing GFP but lacking CD45 (red) (arrowhead). T, tubule. (B) Confocal images of 7-day postischemic kidney from EGFP chimeras labeled with anti-CD31 antibodies (red). Note EGFP-positive cells coexpressing CD31 (arrowheads) in the 2D image. Also note that the endothelial cell nucleus expresses EGFP but not CD31, which is not expressed in nuclei (arrows). (C) Confocal images from 7-day postischemic kidney from EGFP chimeras labeled with antibodies against vWF (red). Note EGFP-positive cell coexpressing vWF in cytoplasmic granules (arrowhead) in the 2D image. Also note that the endothelial cell nucleus expresses EGFP but not vWF, which is not expressed in nuclei (arrows). (D) Quantification of bone marrow–derived cells expressing markers of endothelial cells through determination of the number of peritubular cells coexpressing EGFP and endothelial markers in contralateral (control) and 7-day postischemic kidneys. Data are presented as percent of vWF-positive or CD31-positive cells that express EGFP. Scale bars: 50 μm.
The Y chromosome is found in postischemic kidney peritubular but not tubular cells in sex-mismatched chimeras when assessed by deconvolution microscopy. Female mice with marrow transplants from male mice were subjected to bilateral and unilateral ischemic injury. At both 7 and 15 days after injury, in situ hybridization was used to assess harvested tissue for the presence of Y chromosome–bearing cells. As expected, the majority of cells in the spleen (Figure 5B) and thymus (data not shown) had a nuclear Y chromosome, which confirmed chimerism. In male postischemic kidneys, the Y chromosome was clearly visible by epifluorescent microscopy in 57.6% ± 6.9% of tubular cells (data not shown).
In female mice with male bone marrow, tubular cells are not derived from bone marrow cells following I/R injury to the kidney. (A) Fluorescent image of kidney outer medulla at day 15 following bilateral ischemic injury. The section was hybridized with an FITC-conjugated probe for the Y chromosome and counterstained with lotus lectin (red) highlighting proximal tubular cells and DAPI showing nuclei. Note that many interstitial cells (arrowheads) exhibit the Y chromosome, but none of the regenerated tubular cells stain for the Y chromosome. (B) Section of spleen showing that the majority of nuclei stain positively for the Y chromosome. (C) Detailed view of a proximal tubule, showing a tubular cell nucleus apparently containing a Y chromosome when viewed by epifluorescence. (D) An ultrathin deconvolution image through the same section as shown in C. The fluorescent label can be clearly seen to reside outside of the nucleus. (E and F) Sections hybridized with FITC-conjugated Y chromosome probe were counterstained with anti-vWF antibodies (red). (E) Peritubular capillary showing endothelial cell nucleus with Y chromosome. (F) Section of the image in E obtained by deconvolution microscopy confirming vWF staining and a nucleus containing the Y chromosome all in 1 plane. Scale bars: 50 μm.
Postischemic kidneys from 13 chimeric mice were analyzed at days 7 and 15 by in situ hybridization. In each case, chimerism was confirmed by simultaneous assessment of hematopoietic organs (Figure 5B). To enhance proximal tubular staining, we also counterstained sections with lotus lectin (Figure 5A). We observed occasional Y chromosome–positive tubular cells in the contralateral, sham-operated, and postischemic kidneys by epifluorescence. The frequency of positive tubular cells in the 15-day postischemic kidneys was 3.54 ± 0.59 per 40 HPFs. The frequency was also measured as a percentage of all tubular cells and found to be 0.06% ± 0.01%. This frequency was no different from that of positive tubular cells seen in sham-operated and contralateral kidneys and in kidneys from nonchimeric female mice. The same sections were scrutinized further by deconvolution microscopy. This provided not only exceptional 3D detail of the apparently positive tubular cells, but also a 3D composite for detailed analysis of overlying cells and enhanced ability to determine whether the Y chromosome was within the confines of the nucleus (Figure 5, C and D). We examined 16 apparently positive tubular cells in postischemic kidneys of chimeric mice and 5 kidneys from female, nonchimeric mice. In all but 1 case, the positive tubular cell was clearly an artifact, due to a rare overlying leukocyte or intratubular monocyte or, most frequently, nonspecific aggregates of fluorescent probe (Figure 5, C and D). In the 1 example where tubular cell labeling could not be excluded, it was unclear whether this was a leukocyte or a tubular cell. In the 15-day postischemic kidneys, there was only approximately a 2-fold increase in the level of interstitial leukocytes compared with controls. Since the apparently positive tubule cells were artifacts, sometimes due to leukocytes, one might have expected an increase in the frequency of artifact in postischemic kidneys. However, at such a low frequency, at a time when there was only a 2-fold increase in the level of interstitial leukocytes, this small increase in leukocyte levels was not reflected by an apparent increase in Y chromosome–positive tubular cells in 15-day postischemic compared with sham-operated or contralateral kidneys.
To determine in these chimeras whether any peritubular cells of bone marrow origin might be peritubular endothelial cells, we counterstained hybridized sections with the endothelial marker vWF. As was found in the GFP chimeras, small numbers of endothelial cells had the Y chromosome (Figure 5E). These results were confirmed by deconvolution microscopy (Figure 5F).
Bacteria β-gal is not detected in tubular cells of β-gal chimeras following I/R injury. Prior studies assessing the contribution of bone marrow derived cells to I/R injury and repair have reported high levels of engraftment into tubule cells when β-gal–expressing bone marrow cells were used (15, 16). Using a similar approach, we tested the I/R model at 6 weeks after bone marrow transplantation. We confirmed chimerism of irradiated mice transfused with bone marrow from mice expressing the LacZ gene by staining buffy coat leukocytes with X-gal. Nearly all (91% ± 7%) leukocytes stained blue compared with 0% ± 0% of wild-type C57BL/6J blood leukocytes. In addition, when kidneys were harvested for analysis, bone marrow was also stained with X-gal and was found to be nearly completely reconstituted with X-gal–stained cells (Figure 6, A and B). Tissue sections of hematopoietic organs, spleen, and thymus (n = 5 per time point) from the chimeric mice also exhibited widespread positive staining with X-gal, which confirmed chimerism in the organs of the mice, as did peritoneal macrophages and lung alveolar macrophages (Figure 6C). For example, upon analysis of HPFs, 94% ± 8% of peritoneal cells (Figure 6C) and 79% ± 15% of spleen cells from chimeric mice (n = 10) stained blue with X-gal, whereas 0% ± 0% of peritoneal cells and 0% ± 0% of spleen cells from wild-type C57BL/6J animals stained blue with X-gal (data not shown).
Confirmation of chimerism in mice transplanted with β-gal–expressing bone marrow. (A) Fixed bone marrow cells stained blue with X-gal. (B) Percentage of bone marrow cells that stained blue with X-gal in control C57BL/6J mice and chimeric mice (n = 5 per time point) subjected to I/R followed by 2 days and 7 days of recovery. (C) Peritoneal cells as well as cells of thymus, spleen, and lung from chimeric mice all stained positively with X-gal. Scale bars: 50 μm.
X-gal staining was elevated in ipsilateral and contralateral kidneys of mice at 2, 7 (n = 9), and 21 days (n = 5) following unilateral I/R injury induced 6 weeks after bone marrow transplantation. Viewing HPFs (magnification, ×400), we detected X-gal staining in a small number of tubular cells. These X-gal–positive tubules colabeled with the proximal tubular cell marker gp330 (Figure 7A). This degree of staining, however, did not differ markedly from that seen in the contralateral kidneys and represented far fewer cells (Figure 7B) than would be expected based on the percentage of PCNA-positive cells (Figure 1C).
X-gal stains tubular cells in the kidney of wild-type mice, reflecting endogenous β-gal activity, but can be distinguished from bacterial β-gal when high-pH X-gal staining or anti–β-gal antibodies are used. (A) Occasional tubular cells stained blue, indicating β-gal, 7 days following I/R injury in chimeric mice (arrowheads). X-gal–stained cells colabeled with the proximal tubular cell marker gp330, seen as green fluorescence (inset). (B) The number of X-gal–positive cortical and outer medullary tubular cells per 40 HPF in contralateral kidneys (0 days) and in kidneys of both chimeric and wild-type C57BL/6J mice following I/R injury. Note that I/R resulted in an increase in the number of β-gal–stained cells in both nonchimeric and chimeric mice. (C) Sections at day 7 after I/R labeled with anti–β-gal antibodies. Note the bright staining in spleen cells (left) but no staining in regenerated tubules (right) (D) Sections (magnification, ×100) of I/R kidneys and spleen from wild-type (n = 3 per time point) and chimeric mice (n = 5 per time point) stained with X-gal solution at pH 6.5 or 7.5. Note faint blue tubules in kidneys of wild-type and chimeric animals (compare top and middle panels). Blue staining in kidney tubules but not spleen is suppressed using X-gal in solution at pH 7.5 in chimeric mice (compare bottom and middle panels). (E) Sagittal sections (0.2 mm) of normal C57BL/6J kidney stained with X-gal solution at pH 6.5 (left) and pH 7.5 (right). Note widespread cortical and outer medullary staining at low pH and milder restricted staining in the outer medulla at higher pH. (F) Kidney section (magnification, ×400) from LacZ donor mouse stained with X-gal, pH 7.5. Note intense blue staining of tubules indicative of bacterial β-gal. Scale bars: 50 μm.
Given that the kidney expresses endogenous β-gal (20, 21), the staining in chimeric mice was compared to X-gal staining in kidney sections from nonchimeric wild-type C57BL/6J mice after unilateral I/R injury. Sections taken from these nonchimeric mice also had small numbers of tubular cells that stained blue with X-gal solution prepared as described in Methods (Figure 7B). These data suggested that X-gal staining in chimeras might be detecting endogenous β-gal activity. Frozen sections from 48-hour and 7-day postischemic tissue were immunolabeled with antibacterial β-gal antibody. By this method, spleen cells labeled strongly for β-gal in chimeric mice (Figure 7C), but the antibody did not detect β-gal in the postischemic tubular cells (Figure 7C). To clarify further whether the β-gal activity detected by X-gal staining was endogenous or bacterial, we used a commercially produced β-gal staining set (Roche Diagnostics Corp.) designed to minimize staining from mammalian β-gal. With these reagents, bone marrow, spleen (Figure 7D), and thymic tissue in chimeric mice stained to a level equal to that seen when the X-gal solution prepared in our laboratory was used as a positive control. By contrast, however, when this commercial X-gal solution was used, tubular cells did not stain for β-gal either following I/R injury or in contralateral kidneys in chimeric mice (Figure 7D). Careful review of all the ipsilateral and contralateral kidney sections from chimeric mice, taken 2, 7, and 21 days after ischemia, that were stained by this method revealed only 1 positive tubular cell in a contralateral kidney. Tubular cells did not stain positively for β-gal in control C57BL/6J mice (data not shown).
What might account for occasional positive staining with one X-gal solution and not another? A reduction in pH of the X-gal staining solution has been reported to result in staining of the mammalian form of β-gal (21). Using the commercial X-gal solution, which has a pH of 7.5, we evaluated whether tubules of postischemic nonchimeric mice were more likely to stain positively if we lowered the pH. When pH was lowered to 6.5, we could detect positively stained tubular cells in wild-type C57BL/6J kidney sections (but not spleen sections; Figure 7D). Therefore, when X-gal staining solution has a pH less than 7.5, there is significant staining of endogenous β-gal activity in kidney sections. Careful adjustment of pH can minimize this unwanted staining. These findings are exemplified by the staining of 0.2-mm tissue sections of normal wild-type C57BL/6J kidneys that had been perfusion-fixed with PLP (2% paraformaldehyde [PFA], 75 mM L-lysine, and 10 mM sodium periodate) (Figure 7E). The commercial staining set (pH 7.5; Roche Diagnostics Corp.) detected a minimal level of endogenous β-gal. Much stronger detection was seen when the pH was reduced to 6.5. These images indicate that endogenous β-gal was detected in the kidney tubule at pH 6.5. When detection in thick tissue sections (0.2 mm) was minimized (at pH 7.5), tubule staining was attenuated but could still be seen faintly in the outer medulla, precisely the segment of nephron most susceptible to injury from ischemia (Figure 7E). Thick tissue sections (0.2 mm) of wild-type C57BL/6J kidneys 2 days after ischemia, stained at pH 7.5, also showed a weak diffuse cortical staining pattern, and at pH 6.5, there was much stronger staining in cortical tubules (data not shown). Note that kidney sections from donor LacZ mice had strong kidney tubule staining with X-gal at pH 7.5 (Figure 7F).
After I/R injury, the LacZ chimeric mouse kidneys did not show evidence for endothelial cell replacement, although there were positively stained leukocytes in the interstitium. We reviewed tissue sections from the LacZ donor mice and found that endothelial cells did not stain with the X-gal solution, which suggests that bacterial β-gal was only weakly expressed in the endothelium. Bone marrow–derived endothelial cells would therefore not be expected to be identified in this chimeric mouse.
Intravenous therapy for I/R injury with bone marrow MSCs ameliorates injury without differentiation into renal structures. Although our data indicated that bone marrow stem cells make only a minor direct contribution to replenishment of postischemic renal structures, we wondered whether direct infusion of bone marrow–derived MSCs might promote stem cell differentiation into renal structures, in particular endothelial cells. Bone marrow MSCs have multipotentiality, as evidenced by their ability to differentiate into a variety of mesenchymal cell types (22). We purified MSCs from EGFP mice (23). These cells did not express lineage markers, or c-kit, but expressed CD34 and Sca-1 when assessed by flow cytometry (Figure 8A). Importantly, in addition to c-kit (which is expressed by endothelial cells), they did not express other endothelial cell markers, including CD31. The MSCs were highly fluorescent with EGFP and could be induced in vitro to differentiate into endothelial-like structures and adipocytes (Figure 8, B and C), which confirmed the multilineage potential of the MSCs. In the case of endothelial-like structures, these labeled strongly with anti-CD31 antibody (Figure 8B). As quantitated by flow cytometry, 1.8% ± 0.5% of MSCs exhibited a 100- to 1,000-fold increase in expression of CD31 when cultured on Matrigel (data not shown), which provides further evidence that MSCs have the capacity to develop an endothelial phenotype. MSCs were cultured on tissue culture grade plastic. Mice were subjected to bilateral I/R injury, followed by i.v. injections of MSCs (0.5 × 106 cells per injection) immediately after surgery and at 24 hours after ischemia. In our initial studies, peak plasma creatinine levels at 24 hours were no different in mice receiving MSCs or an injection of control medium (PBS) (Figure 8E). Organs examined by fluorescent microscopy at 24 and 48 hours provided no evidence for implantation of MSCs. The experiments were repeated using MSCs that had been cultured on Matrigel. A control cell type, embryonic fibroblasts, was prepared identically. Injection of MSCs reduced the severity of ARF as measured by the creatinine level at 24 hours (Figure 8E). This effect was, however, also seen when embryonic fibroblasts, grown on Matrigel, were injected (Figure 8F). We examined these kidneys by fluorescence microscopy 24 hours, 48 hours, 7 days, and 15 days after I/R injury. There was no evidence of MSCs in the kidney parenchyma. We did not find 1 example of EGFP-expressing cells in any of these kidneys. Furthermore, we examined other organs, in particular lung and spleen, where there was also no evidence for MSCs implantation. Analysis of bone marrow at 15 days by flow cytometry revealed a small population of EGFP-fluorescent cells (less than 1%), which suggests that MSCs had homed to the marrow cavities.
Bone marrow MSCs have multilineage potential and are protective in ischemic injury without differentiating into tubule cells. (A) Flow cytometry of MSCs for cell surface stem cell markers. Compared with isotype control labeling, which was used to define the marker (data not shown), MSCs lacked expression of c-kit and CD31 but expressed CD34 and Sca-1 (values are expressed as percent positive compared with isotype control antibody labeling). (B) Photomicrograph of MSCs differentiated into capillary-like structures (left). The same structure shows strong immunofluorescent labeling with the endothelial marker CD31 (right). (C) Photomicrograph of MSCs differentiated in vitro into adipocytes and labeled with oil red O for lipid. (D) Fluorescence micrograph showing an EGFP-expressing MSC in the cortex of the postischemic kidney 2 hours after intraparenchymal injection of EGFP-labeled MSCs. (E) Plasma creatinine levels 24 hours after 30-minute bilateral I/R renal injury followed by i.v. injection of control PBS or 0.5 × 106 MSCs cultured on plastic (n = 4 per group). (F) Plasma creatinine levels 24 hours after 30-minute bilateral I/R renal injury followed by i.v. injection of control PBS, 0.5 × 106 MSCs cultured on Matrigel, or embryonic fibroblasts (Fibro) cultured on the same matrix. Note that the level of creatinine was significantly higher in PBS-treated mice (n = 7 per group; **P < 0.01, ANOVA). Scale bars: 50 μm.
We further explored the potential of MSCs to become renal structures by direct intrarenal parenchymal injection of MSCs following ischemic injury. Two hours following injection of MSCs (0.5 × 106) into the cortex of the normal or postischemic kidney, viable MSCs could be identified by microscopy (Figure 8D). However, 24 hours later, there was no evidence of EGFP cells in the kidney. These results are comparable to those of an earlier study, in which the fate of MSCs was traced in irradiated mice (22). In this study, MSCs homed to the bone marrow only. Our data indicate that, while an injection of cells i.v. can have a therapeutic effect on the course of functional impairment due to I/R when cells are grown under specific conditions, this effect is independent of implantation and differentiation into renal structures.
Hematopoietic lineage–negative leukocytes. Although we found no evidence for differentiation of bone marrow–derived cells into tubular cells, we did find evidence for differentiation into endothelial cells in the injured kidney. It was important therefore to determine whether renal injury mobilized stem cells from the bone marrow to the blood, as has been suggested in other reports (15, 24). Bone marrow–derived HSCs and MSCs lack lineage markers, but the identity of cell surface markers of endothelial precursors remains ambiguous. In some studies, endothelial precursors have also been shown to lack lineage markers (24, 25). Blood leukocytes purified from healthy mice or mice 24 and 48 hours following ischemic injury were labeled with antibodies to lineage markers and analyzed by flow cytometry (Figure 9). In healthy mice, a small population of leukocytes (less than 1%) was hematopoietic lineage–negative (lin–). We found no evidence for an increase in this negative population at 24 hours (data not shown) or at 48 hours (Figure 9) following injury.
The proportion of lin– blood leukocytes does not increase following I/R injury. Flow cytometry plots (representative of n = 6) of blood leukocytes from healthy mice (left) or mice 2 days following bilateral renal ischemia (right), labeled with PE-fluorescent antibodies against lineage markers (Lin-PE) (CD11b, Gr-1, CD4, CD8, B220, Ter-119) or IgG control antibodies. Note that a minority (<1%) of leukocytes were lin– in control mice, and this percentage was not increased in mice 2 days following I/R injury.