Clinical Research

APOL1 Localization in Normal Kidney and Nondiabetic Kidney Disease

Sethu M. Madhavan, John F. O'Toole, Martha Konieczkowski, Santhi Ganesan, Leslie A. Bruggeman and John R. Sedor
JASN November 2011, 22 (11) 2119-2128; DOI: https://doi.org/10.1681/ASN.2011010069

Abstract

In patients of African ancestry, genetic variants in APOL1, which encodes apolipoprotein L1, associate with the nondiabetic kidney diseases, focal segmental glomerulosclerosis (FSGS), HIV-associated nephropathy (HIVAN), and hypertensive nephropathy. Understanding the renal localization of APOL1 may provide clues that will ultimately help elucidate the mechanisms by which APOL1 variants promote nephropathy. Here, we used immunohistology to examine APOL1 localization in normal human kidney sections and in biopsies demonstrating either FSGS (n = 8) or HIVAN (n = 2). Within normal glomeruli, APOL1 only localized to podocytes. Compared with normal glomeruli, fewer cells stained for APOL1 in FSGS and HIVAN glomeruli, even when expression of the podocyte markers GLEPP1 and synaptopodin appeared normal. APOL1 localized to proximal tubular epithelia in normal kidneys, FSGS, and HIVAN. We detected APOL1 in the arteriolar endothelium of normal and diseased kidney sections. Unexpectedly, in both FSGS and HIVAN but not normal kidneys, the media of medium artery and arterioles contained a subset of α-smooth muscle actin-positive cells that stained for APOL1. Comparing the renal distribution of APOL1 in nondiabetic kidney disease to normal kidney suggests that a previously unrecognized arteriopathy may contribute to disease pathogenesis in patients of African ancestry.

Two genome-wide admixture scans identified a highly significant association on chromosome 22q12 between nondiabetic kidney disease and African ancestry.1,2 The associated region contains many genes and initial investigations focused on nonmuscle myosin heavy chain IIA, encoded by MYH9, due to its podocyte expression and association with rare Mendelian giant platelet disorders that are also characterized by variably penetrant glomerular diseases.3,4 However, attempts to identify the MYH9 causal variant underlying this association were unsucessful.5,6 Using 1000 Genomes Project sequence data,7 two groups demonstrated that coding variants within the neighboring APOL1 gene,8,9 which is centromeric to MYH9 on 22q12.3, accounted for the association signal. APOL1 is one of six closely related apolipoprotein L family member genes clustered on chromosome 22. APOL1 is restricted to the genomes of humans and some nonhuman primates.10 Circulating APOL1 associates with HDL3 particles and functions as a trypanolytic factor in human serum.11,12 The parasite endocytoses APOL1-containing HDL particles, and once internalized, APOL1 is targeted to the lysosome, where its colicin-like pore-forming activity causes osmotic swelling of the lysosome and trypanosome death. Trypanosome species that infect humans and cause disease have adapted to inhibit APOL1-mediated trypanolysis. Variant APOL1, which encodes the kidney disease risk variants, can kill disease-causing trypanosomes by circumventing the parasite's mechanism to evade lysis. The parasite-killing effect is dominant, requiring a single copy of the risk-variant APOL1 gene, whereas association with kidney disease is best fit by a recessive model.9 Since resistance to trypanosomal infection is a selective advantage in endemic regions, the kidney disease risk variants of APOL1 have been maintained in African populations. Consistent with this premise, the region of chromosome 22 that contains APOL1 shows evidence for positive selection.13 Similar to sickle cell disease, the heterozygous state for APOL1 kidney-disease-associated variants is advantageous, but the homozygous state can result in disease.9

APOL1 is a 43 kD protein that is the only APOL family member with a secreted isoform. Since APOL1 transcripts are expressed in a number of tissues, including the kidney,14,15 we have characterized cellular localization of APOL1 in normal human kidney sections and biopsies from patients with focal segmental glomerulosclerosis (FSGS) and HIV-associated nephropathy (HIVAN) as a first step in understanding the association between APOL1 genetic variation and nondiabetic kidney disease.

RESULTS

APOL1 Localizes to the Glomerulus, Cortical Tubules, and Vascular Endothelium of the Normal Kidney

We verified the specificity of two commercially available antibodies against APOL1 using multiple approaches (Supplementary Figures 1, 2, and 3) and examined formalin-fixed, paraffin-embedded human kidney sections without renal pathology to determine the normal cellular distribution of APOL1 in renal parenchyma. Using immunohistochemistry, we found that all APOL1 immunoreactivity at low magnification was in the renal cortex of normal human kidneys (Figure 1A) relative to control sections, where only faint background resulting from endogenous peroxidase activity was detected (Figure 1 B). Higher magnification views revealed robust staining in a subset of cortical tubules that appeared to be proximal tubules by morphology (Figure 1 C). The localization of APOL1 within proximal tubules was confirmed using the lectin lotus tetragonolobus (Supplementary Figure 4). APOL1-postive tubules were distinct from tubules positive for uromodulin, a marker of thick ascending limb of Henle. Less intense glomerular staining was observed in a subset of cells (Figure 1 C), which was not evident in control sections (Figure 1 D). High magnification views of the glomerulus demonstrated peroxidase staining of cells that are consistent with podocytes with respect to anatomical position and morphology (Figure 1 E). At similar magnification, glomerular cells in control sections showed no immunostaining (Figure 1 F). In addition, signal was detected in the endothelium of small arteries (Figure 1 G), which was not seen in control sections (Figure 1 H).

Figure 1.
Figure 1.

APOL1 localization in the normal adult human kidney using immunoperoxidase staining. Serial, formalin-fixed, paraffin-embedded sections of normal human kidney specimens were immunostained with rabbit polyclonal APOL1 antibody (A, C, E, & G) or purified rabbit polyclonal IgG as control (B, D, F, & H). (A) Low magnification image showing predominant APOL1 staining in the renal cortex, with relative sparing of medulla. (C) APOL1 expression is seen in a glomerulus and some cortical tubules. (E) Magnified image from boxed region in (C) shows APOL1 staining in a cell with anatomical position and morphologic characteristics of a podocyte, abutting the outer lamina of the glomerular basement membrane with red blood cells (asterisk) contained in a capillary loop. (G) APOL1 staining is also seen in extraglomerular arterial endothelium (arrow). Comparable images of normal kidney sections (B, D, F, & H) incubated with nonimmune purified rabbit polyclonal IgG have minimal staining consistent with residual endogenous peroxidase activity. Scale bars: 300 μm (A & B), 50 μm (C, D, G, & H).

APOL1 Colocalizes with Podocyte Markers in the Normal Glomerulus

We used immunofluorescent confocal microscopy to colocalize APOL1 with markers for podocytes and mesangial and endothelial cells in normal human kidney sections (Figure 2). The cytosol of cells within glomeruli staining for APOL1 was completely outlined by GLEPP1, a podocyte transmembrane receptor phosphatase (Figure 2 A–F), suggesting localization of APOL1 within the podocyte. This pattern was consistent in additional fields stained for GLEPP1 and also when antibodies to another podocyte marker (synaptopodin) were used (Supplementary Figure 5). We were unable to detect significant glomerular expression of α-smooth muscle actin (α-SMA) in the mesangium of the normal kidney. Mural cells in arterial vessels near the vascular pole of the glomeruli did express α-SMA, which did not overlap with APOL1 staining in cells that are endothelia by their location (Figure 2 G–I). Staining with antibodies against APOL1 and CD31, an endothelial marker, did not overlap in the glomerular capillaries, with the CD31 signal clearly on the opposite side of the glomerular basement membrane from the APOL1 signal (Figure 2 J–L). In contrast, CD31 and APOL1 antibodies identified the same cells in arterial vessels.

Figure 2.
Figure 2.

Colocalization of APOL1 with cell markers in the normal adult human kidney. Confocal immunofluorescent microscopy demonstrates (A, D, G, & J, green) APOL1 staining in the glomerulus of the normal kidney. (B & E, red) GLEPP1, a podocyte transmembrane protein tyrosine phosphatase, demarcates podocyte plasma membranes. (C & F) Merged images show that the APOL1 signal (green) is completely circumscribed by GLEPP1 staining (red) and appears to be present in the same cells. (D–F) Magnified view of the indicated region in (C) shows that APOL1 (green) localizes to two cell bodies (arrowheads) outlined by GLEPP1 staining (red), and nucleated endothelial cells lining the adjacent capillary loop (asterisks) show no APOL1 staining. (H) Cells within normal adult human glomeruli do not express α-SMA (red); however, positive α-SMA staining is seen in an adjacent arterial wall (double arrows). (I) Merged image shows APOL1 staining (green) in the adjacent arterial endothelium (arrow) but not within the arterial wall. (K & L, red) CD31 staining of the glomerular endothelium does not colocalize with APOL1 (green). The inset in (L) is a magnified image of the indicated region in that panel. (C, F, I, & L, blue) Nuclei were visualized with TOTO-3 staining. Scale bars: 50 μm (A–C & G–L), 5 μm (D–E).

APOL1 Risk–variant Genotypes of FSGS and HIVAN Biopsy Samples

We obtained deidentified, archival renal biopsy specimens demonstrating FSGS (n = 8) or HIVAN (n = 2), along with patient ages, genders, and self-reported races, and determined the genotype at the G1 and G2 loci with Sanger sequencing of DNA extracted from an additional section cut from tissue blocks9 (Table 1). The risk genotype was successfully determined in nine of the 10 samples examined in this study; Supplementary Figure 6 shows the sequencing data from each patient. Although the G1 risk variant is more common than the G2 risk variant, it was not found in the biopsies examined. The G2 risk allele was homozygous in two African American patients and was heterozygous in one African American and one Caucasian patient, likely reflecting African admixture in this individual. Specimens were reviewed to confirm that pathology was characteristic of FSGS and HIVAN, and images of representative renal pathology specimens, using standard histochemical stains, are shown as supplementary data (Supplementary Figure 7).

View this table:
Table 1.

APOL1 risk genotypes in FSGS and HIVAN samples

APOL1 Is Increased in the Vasculature and Decreased in Podocytes in FSGS

Since genetic variants in APOL1 have been associated with nondiabetic kidney diseases in African Americans, we determined if the renal distribution of APOL1 varied with respect to risk genotype in the biopsies. FSGS samples were examined for glomerular APOL1 expression using immunohistochemistry (Figure 3 A–C). The overall immunohistochemical staining for APOL1 appeared to be reduced in the FSGS biopsy specimens with fewer APOL1-positive tubules as seen in the low power view (Figure 3A). The number of cells staining for APOL1 within the FSGS glomeruli appeared to be less numerous compared with normal glomeruli (Figures 3B, 1C). In contrast to normal kidneys, a robust signal was detected in vessel walls at the glomerular vascular pole (Figure 3 A,B) and small arteries in the renal parenchyma (Figure 3C), which had not been observed in normal kidney sections (Figure 1G).

Figure 3.
Figure 3.

Altered renal localization of APOL1 in focal segmental glomerulosclerosis (FSGS). Immunoperoxidase (A–C) and confocal immunofluorescent images (D–L) of kidney biopsy specimens from patients with FSGS stained using anti-APOL1 antibody. (A) Reduced APOL1 staining is seen in glomeruli and cortical tubules compared with the normal kidney. In contrast, glomerular vascular tuft demonstrates prominent APOL1 expression (arrowheads). (B) Selected glomerulus from (A) shows diminished APOL1 staining (arrows); however, the juxtaglomerular arteriole wall is prominently APOL1 positive (arrowhead). (C) In contrast to the normal kidney, APOL1 also appears in the vessel wall of the renal arterioles. (D & F, green) APOL1 signal is diminished in the glomerulus, but glomerular GLEPP1 expression (E & F, red) is relatively preserved in early FSGS. The merged images (F) highlight these differences, where all APOL1 (green) positive cells stain for GLEPP1 (red) but not all GLEPP1 expressing cells stain for APOL1. (G–I) Residual glomerular APOL1 signal (G & I, green) in early FSGS does not colocalize with the α-SMA staining (H & I, red) of reactive mesangium. However, APOL1 signal (G & I, green) is observed in the wall of the arteriole at the vascular pole of the glomerulus (asterisk) and colocalizes with α-SMA (H & I, red), which labels vascular smooth muscle cells in the extraglomerular arteriolar wall (asterisk). (J–L, green) APOL1 remains diminished in early FSGS and does not colocalize with glomerular anti-CD31 staining (K & L, red), which identifies endothelial cells, but does colocalize in the extraglomerular vascular endothelium (arrowheads). (L) Merged image shows overlap of APOL1 (green) and CD31 (red) staining in the extraglomerular vascular endothelium (arrow) but not within the glomerulus. (F, I, & L, blue) Nuclei were visualized with TOTO-3 staining. Scale bars: 100 μm (A), 50 μm (B–L).

To further characterize changes in APOL1 glomerular distribution of FSGS samples, we used the same cell lineage markers described above to examine mildly affected (Figure 3 D–L) and severely affected glomeruli (Supplementary Figure 8). The overall fluorescence intensity of APOL1 signal in the glomeruli of FSGS and HIVAN samples was diminished relative to glomeruli from samples without renal pathology (Supplementary Figure 9). Residual APOL1 signal remained colocalized with GLEPP1 but some podocytes that still expressed GLEPP1 has diminished or absent APOL1 expression (Figure 3 D–F). Glomerular staining of α-SMA in FSGS samples was increased, labeling reactive mesangium, but did not colocalize with APOL1 (Figure 3 G–I). However, APOL1 did colocalize with α-SMA staining in the vascular smooth muscle cells of arterioles at the glomerular vascular pole (Figure 3 G–I). CD31 staining of the glomerular endothelium did not colocalize with APOL1 (Figure 3 J–L). Interestingly, the FSGS samples demonstrated a striking increase of APOL1 in the vascular wall of the small renal arteries outside glomeruli, which was not found in the normal kidney (Figure 4). As in the vessel wall of the glomerular vascular pole (Figure 3 G–I), APOL1 colocalized with α-SMA staining in some arterial wall cells (Figure 4). Endothelial cells continued to stain for APOL1 in FSGS biopsy specimens (Figure 4). APOL1 staining patterns were consistent in all biopsies, regardless of APOL1 G2 genotypes.

Figure 4.
Figure 4.

De novo localization of APOL1 to the renal arterial wall in FSGS. Confocal immunofluorescence imaging of medium sized renal arterioles from the normal human kidney (A–C) and from FSGS (D–F). (A & C, green) APOL1 signal is identified in cells anatomically consistent with endothelium of medium-sized renal arterioles (arrowheads) in the normal adult human kidney and adjacent tubular segments but does not colocalize with vascular smooth muscle cells of renal arterioles stained with anti-α-SMA antibody (B & C, red). APOL1-positive cells are located in the luminal wall of the blood vessel with surrounding α-SMA positive vascular smooth muscle cells, consistent with endothelial localization. (D & F, green) Representative cross-section of a renal arteriole from an FSGS biopsy demonstrates persistent endothelial APOL1 signal and de novo appearance within the vessel wall compared with the normal arteriole (A). (E & F, red) Anti-α-SMA antibody staining identifies APOL1-positive cells as vascular smooth muscle (F, green). The vascular endothelium remains positive for APOL1 staining (arrowhead). (C & F, blue) Nuclei were visualized with TOTO-3 staining. Scale bars: 50 μm (A–F).

Alterations in APOL1 Localization Are Consistent between FSGS and HIVAN

While only two HIVAN samples were available for evaluation, overall the alterations in APOL1 distribution were similar to those observed in the FSGS samples. There appeared to be fewer APOL1 positive tubules (Figure 5A), less glomerular signal (Figure 5 B), and de novo appearance in the vessel wall of renal arteries (Figure 5 C). When present in glomeruli, APOL1 remained colocalized with podocyte markers (Figure 5 D–F) but not with the endothelial marker CD31 (Figure 5 H–J). α-SMA and APOL1 did not colocalize within the glomeruli (Supplementary Figure 10 A–C) but, similar to the FSGS biopsies, did colocalize within the vascular smooth muscle cells of small renal arteries (Supplementary Figure 10 D–F). APOL1 localization within the endothelium of renal arteries in HIVAN biopsies (Supplementary Figure 10 D–F) appeared similar in intensity and distribution to that identified in biopsies of FSGS and normal sections (Figure 4).

Figure 5.
Figure 5.

APOL1 redistribution is consistent between FSGS and HIVAN samples. Immunoperoxidase (A–C) and confocal immunofluorescent images (D–I) of kidney biopsy specimens from patients with HIVAN stained using anti-APOL1 antibody. (A) Reduced APOL1 staining in glomeruli and cortical tubules compared with the normal kidney. (B) Selected glomerulus from (A) shows diminished APOL1 staining (arrowhead). (C) In contrast to the normal kidney, APOL1 also appears in vessel wall of the renal arterioles. (D & F, green) APOL1 signal is diminished in the glomerulus, but remains colocalized with GLEPP1 expression (E & F, red), which is also diminished in HIVAN. (H & J, green) Glomerular APOL1 signal does not overlap with CD31 staining (I & J, red), which identifies glomerular capillary endothelium. (F & I, blue) Nuclei were visualized with TOTO-3 staining. Scale bars: 100 μm (A), 50 μm (B–I).

APOL1 Is Expressed in Renal Cell Lines and Induced by Inflammatory Mediators

To examine whether APOL1 is endogenously expressed in the kidney or absorbed from the circulation, we prepared lysates from human podocyte, proximal tubular, and vascular endothelial cell lines grown in culture. Lysates were immunoblotted for APOL1 and a single band was detected at the expected molecular weight (Figure 6A). RT-PCR was performed on RNA extracted from a human podocyte cell line and the transcript of expected size was detected (Figure 6 B). Based on prior reports, APOL1 expression can be induced by inflammatory mediators such as recombinant tumor necrosis factor (TNF) and lipopolysaccharide (LPS). Stimulation of human microvascular endothelial cells with TNF or LPS induced expression of APOL1 relative to tubulin (Figure 6 C–D).

Figure 6.
Figure 6.

APOL1 is expressed in cultured human kidney cells and is induced in vascular endothelial cells with TNF/LPS treatment. (A) APOL1 is detectable by immunoblotting in cultured human podocytes, proximal tubule (HPRT) cells, and CHO cells transfected with an APOL1 expression construct. The blot is representative of three independent experiments. (B) APOL1 transcript is detectable in total RNA isolated from cultured human podocytes. The blot is representative of three independent experiments. (C, D) In vitro expression of APOL1 was detected in unstimulated and TNF- or LPS-stimulated human microvascular endothelial cells (hMVECs) by immunoblotting and normalized by the expression of tubulin (C). APOL1 expression was quantified by densitometry in unstimulated hMVECs and hMVECs stimulated with TNF or LPS (n = 3) (D). Results were normalized to tubulin expression and expressed as the mean ± SE; p values were calculated with ANOVA, followed by t test, to make intergroup comparisons. *P < 0.05.

DISCUSSION

We have characterized the cellular localization of APOL1 in sections from a single healthy kidney and from biopsies demonstrating FSGS and HIVAN. The key results demonstrate that, in the absence of pathology, APOL1 is detected in podocytes, proximal tubules, and in medium-sized arterial and arteriolar endothelial cells. In HIVAN and FSGS, the cellular distribution pattern evolves. The number of APOL1-positive podocytes is diminished before a decrease of the podocyte differentiation markers GLEPP1 and synaptopodin. Tubular APOL1 also diminishes and there is a de novo appearance of APOL1 within cells of the arterial medial wall.

An immediate question is whether renal localization of APOL1 reflects de novo synthesis or uptake from circulation. APOL1 is a component of HDL3 particles and its presence within kidney cells could reflect uptake of this subset of HDL particles. Proximal tubules express CD36, which binds unmodified HDL,16 and the punctate pattern of APOL1 staining in proximal tubules may indicate uptake by an endocytic process. Endothelial cells take-up HDL through scavenger receptor B1,17 and the dyslipidemia associated with nephrotic syndrome is associated with elevated vascular deposition of various lipoprotein complexes.18 In advanced chronic kidney disease, however, HDL production is typically repressed and thus the dynamics of renal exposure to circulating APOL1-containing HDLs during the course of the disease may be complex, especially in kidney diseases caused by infection such as HIVAN.

We were surprised that APOL1 had such robust expression within specific kidney domains and hypothesized that APOL1 may be endogenously synthesized by renal tissue. Although originally cloned as a pancreas-specific apolipoprotein,19 subsequent publications showed APOL1 transcripts in a number of tissues, including in the kidney, consistent with intrarenal APOL1 synthesis.14,15 Our studies have not conclusively established whether the APOL1 detected in the various renal parenchymal cells was synthesized de novo or was produced elsewhere and taken up by renal cells. However, based on our observations in cultured cells and in conjunction with published expression array studies, APOL1 synthesis within the kidney is possible. Both de novo expression and extracellular uptake may uniquely or synergistically contribute to pathogenesis.

Another important point to consider in the context of renal APOL1 expression is its intracellular trafficking and secretion. Although all APOL1 isoforms have signal peptides, regulatory mechanisms of APOL1 synthesis and intracellular trafficking have not been characterized in metazoa. An obvious approach to define the relative importance of circulating and renal-expressed APOL1 in the pathogenesis of nondiabetic kidney diseases would be the examination of transplant registries to determine if APOL1 risk variants in the recipient and/or donor organ confer increased susceptibility to graft loss after transplantation through recurrent or de novo pathologies associated with APOL1 risk variants.

The presence of APOL1 in the normal kidney suggests that it is serving a functional role in the normal kidney. As a constituent of HDL3 particles, it has been assumed, but not demonstrated, that APOL1 participates in cholesterol metabolism. In addition, APOL1 forms ion channels in lipid bilayers20,21 and may disrupt membranes,22 but these studies, while informative about APOL1's trypanolytic activity, do not illuminate function within normal cells. Currently, the function of APOL1 in any of the resident cells of the kidney is entirely speculative. Although we detected a clear change from the normal distribution of APOL1 in both FSGS and HIVAN biopsies, the patterns did not correlate with kidney-disease-associated genotypes. Similar cellular distribution changes were demonstrated in patients homozygous, heterozygous, or null for the G2 risk variant, suggesting that the kidney-disease-associated variants do not cause disease by altering APOL1 trafficking or protein levels. Rather, the G2 risk variant of APOL1 likely results in dysregulation of APOL1's functional role within the kidney cells where it is localized. None of our patients had the G1 risk variants, so we cannot comment on the effect of this variant on APOL1 localization in the kidney. Our study has limitations. Biopsy sample size is small, and population frequencies for the G1 and G2 APOL1 alleles cannot be accurately quantified from nine patient samples. Although technical difficulties in genotyping from biopsies may account for failure to identify patients with the G1 variant, our sequence chromatograms were of high quality (Supplementary Figure 6).

The presence of APOL1 in the podocyte within the glomerular compartment is consistent with the prevailing notion that the podocyte plays a central role in the pathogenesis of FSGS and HIVAN. Since APOL1 is lost from podocytes before GLEPP1 and synaptopodin in FSGS, we reason that its disappearance is not a consequence of generalized podocyte dedifferentiation. Rather, we suspect that the loss of APOL1 from podocytes is either an early marker of dedifferentiation or that preserved APOL1 function is required to maintain a differentiated podocyte phenotype. In vitro work in other cell types has shown that overexpressed APOL1 sequesters phosphatidic acid and cardiolipin and promotes autophagocytic cell death.23,24 However, these observations on APOL1 overexpression do not readily provide insight into APOL1-regulated kidney disease pathogenesis. HIVAN and FSGS biopsy findings do not demonstrate marked changes in APOL1 abundances within individual podocytes compared with normal kidney sections, although fewer cells express APOL1 within the disease glomeruli. If APOL1, at normal expression levels, does regulate autophagy, variant APOL1 may cause disease by dysregulating autophagic pathways, similar to studies in mice with podocyte deletion of autophagy-related 5 (ATG5) caused glomerulopathy in aging mice.25

The vascular distribution of APOL1 was restricted to the endothelium of arterioles in normal kidney sections, and we were unable to detect APOL1 in the glomerular capillary endothelium of normal or diseased kidney sections. Previous reports have shown in situ expression of APOL1 and APOL3 transcripts in aortic endothelial cells.26,27 The endothelial localization of APOL1 in the arteriolar endothelium remained in the FSGS and HIVAN samples, and this expression may be increased in disease. With a role in combating Trypanosome infection, APOL1 is considered an acute phase reactant, and like other acute phase reactants, its production is regulated by cytokines that mediate immune responses such as TNF and type I interferons.26,27 Our in vitro studies similarly found that vascular endothelial cells induced APOL1 expression in response to TNF and bacterial endotoxin, consistent with prior observations with aortic endothelia cells.26,27

The de novo appearance of APOL1 in the arterial vessel wall was unexpected. In contrast to the normal kidney, APOL1 was dramatically increased in the vascular smooth muscle cells of the medium size and arteriolar vessel walls in the FSGS and HIVAN biopsies. The origin of these APOL1-positive cells is not clear but the partial colocalization of APOL1 and α-SMA suggests the cell is of vascular smooth muscle lineage. Arterial vascular wall remodeling is a characteristic of many disease processes, and the vascular smooth muscle cell exhibits phenotypic plasticity.28 Alternatively, APOL1-positive cells could reflect migration of activated adventitial cells into the medial layer of the arteriole.29 Prior reports have demonstrated that vascular sclerosis is worse in biopsies from African American compared with Caucasian30,31 patients, suggesting the hypothesis that the de novo appearance of variant APOL1 in small renal arteries contributes to kidney disease pathogenesis.

Although our conclusions should be tempered by the small sample size, we have shown that APOL1 is present in the normal kidney and may be endogenously expressed, and that the cell compartments in which it appears change in HIVAN and FSGS. The cellular distribution patterns do not correlate with kidney-disease-associated APOL1 variants. While the podocyte is an attractive focus for further work, given the evidence for podocyte dysfunction in glomerular disease, the abundant APOL1 expression in cortical tubules may contribute to the pathogenesis of the tubulointerstitial disease that characterizes both HIVAN and FSGS. The de novo appearance of APOL1 in renal arteries and arterioles is intriguing and suggests that vascular wall remodeling may also contribute to FSGS and HIVAN pathogenesis.

CONCISE METHODS

Patient Samples

We obtained formalin-fixed, paraffin-embedded normal human kidney sections (Imgenex Corp, San Diego, CA) and deidentified, archival, formalin-fixed, paraffin-embedded human kidney biopsy specimens assigned a histopathologic diagnosis of FSGS or HIVAN by a clinical pathologist. Age, self-reported race, gender, and histopathologic diagnosis were obtained for each biopsy sample. Samples were otherwise deidentified. All studies involving human tissues were performed with approval of the institutional review board of the MetroHealth System campus, Case Western Reserve University. Eight samples with FSGS and two samples of HIVAN were obtained for these studies.

Cell Lines

A conditionally immortalized, human podocyte cell line has been described previously.32 Human proximal tubule cells immortalized by adeno-12/SV40 transformation were obtained from the laboratory of Dr. Jeffrey Schelling and were cultured in DMEM:F12 1:1 with 10% FBS and 1% PCN/Strep at 37 °C in 5% CO2. Primary human microvascular endothelial cells from lung (hMVECs; Lonza Walkersville, Inc., Walkersville, MD) were cultured in EBM-2 basal media with EGM-2 MV supplements (Lonza Walkersville, Inc.) at 37 °C with 5% CO2 and used at the seventh passage number. For experiments examining the stimulation of APOL1 expression in hMVECs, cells were grown in culture for 48 h to confluence, then treated for an additional 24 h before protein extraction with recombinant human TNF (R&D Systems, Minneapolis, MN) at [20 ng/ml], with LPS from the O55:B5 strain of Escherichia coli (Sigma) at [1 ng/ml], or no additions as an unstimulated control.

Reagents and Anti-APOL1 Characterization

Reagents used in these studies were obtained (Fisher Scientific, Pittsburgh, PA) unless otherwise stated. The specificity of two commercially available anti-APOL1 antibodies was characterized in cell culture and kidney tissue (Supplementary Figures 1–3). We found the polyclonal anti-APOL1 antibody, HPA018885 (Sigma, St. Louis, MO), raised in rabbit using a recombinant fusion protein containing amino acids 263 to 398 of human APOL1 isoform 1, NM_003661.3, to be superior to the polyclonal anti-APOL1 antibody (Abcam, Cambridge, MA) raised in goat using a synthetic peptide corresponding to amino acids 386 to 397 of human APOL1 isoform 1. The rabbit polyclonal APOL1 antibody (Sigma) was used throughout these studies and specifically recognized APOL1 but not APOL3 (Supplementary Figure 1). We did not determine cross-reactivity with other APOL family members, which have similarity to APOL1. Mouse does not express an APOL1 orthologue, and kidney sections incubated with the rabbit anti-APOL1 antibody showed no immunoreactivity (Supplementary Figure 2). In contrast, the goat APOL1 antibody demonstrated significant, nonspecific immunostaining. Immunoreactivity in human kidney sections was abrogated by preincubating the rabbit anti-APOL1 with recombinant, HIS-tagged APOL1, generated as previously published33 (Supplementary Figure 3).

Immunohistochemistry

Formalin-fixed, paraffin-embedded human kidney sections of 4-μm thickness on standard microscopy slides were heated overnight at 40 °C and then at 62 °C for 1 hr. Paraffin was cleared with xylenes, and the section was rehydrated in a graded ethanol series and immersed in ddH2O. Slides were transferred to a pressure boiler containing boiling antigen retrieval solution (10 mM trisodium citrate dihydrate, pH = 6.0, 0.05% tween-20), sealed and heated an additional 4.5 min. Slides were cooled to 25 °C over 1.5 h, rinsed in PBS, pH = 7.4, and washed in PBS, pH = 7.4, with 0.2% tween-20 (PBST). Endogenous peroxidase activity was blocked for 30 min at room temperature with 3% H2O2 in PBS, pH = 7.4, then washed in PBST and blocked for 1 hr at room temperature in 5% serum in PBST. Sections were then incubated overnight at 4 °C in 1% serum, PBS with a polyclonal rabbit anti-APOL1 primary antibody (Sigma, St. Louis, MO) at 1:500 or equivalent purified rabbit polyclonal IgG (Millipore, Billerica, MA). An anti-rabbit immunoperoxidase labeling system, the Vectastain ABC kit, was used according to manufacturer's instructions (Vector Laboratories, Burlingame, CA), washed in PBST, and immersed in PBS. Peroxidase activity was monitored after the addition of substrate using a DAB kit (Vector Laboratories) according to manufacturer's instructions. Sections were counterstained with Hematoxylin, rinsed in tap H20, dehydrated in a graded ethanol series then xylenes, and mounted with permount (Fisher Scientific, Pittsburg, PA). Sections were viewed using an Olympus Bx51 microscope and images captured with Olympus DP71 camera with DP controller software (Olympus America, Inc., Center Valley, PA). Images were prepared for publication using Adobe CS4 software (Adobe Systems, Inc., San Jose, CA).

Immunofluorescence Microscopy

Formalin-fixed, paraffin-embedded sections were prepared as described for immunohistochemistry through the antigen-retrieval protocol. Then the sections were washed in PBST, blocked with 5% serum in PBST, and incubated overnight at 4 °C in PBS, pH = 7.4, 1% serum, 0.1% tween-20 with appropriate primary antibodies, normal purified rabbit polyclonal IgG, or exclusion of primary antibody. APOL1 was detected using a polyclonal rabbit antibody (Sigma, St. Louis, MO) at 1:50. Podocytes were labeled with a mouse monoclonal anti-GLEPP1 antibody (a gift from Dr. Roger Wiggins) at 1:50 or a mouse monoclonal anti-synaptopodin antibody, clone G1D4 (Meridian Life Science, Inc., Saco, ME) at 1:5. Endothelial cells were labeled with a mouse monoclonal anti-CD31 antibody, clone JC70A (Dako, Carpinteria, CA) at 1:20. Smooth muscle was labeled with a mouse monoclonal α-SMA antibody, clone 1A4 (Sigma, St. Louis, MO) at 1:300. After incubation with primary antibody or normal purified rabbit polyclonal IgG sections were washed with PBST, incubated with species specific Fluorophore conjugated secondary antibody (Molecular Probes, Eugene, OR) at 1:400 in PBS, and washed with PBST. Cell nuclei were visualized with TOTO-3 iodide (Molecular Probes, Eugene, OR) staining. Sections were mounted with Vectashield (Vector Laboratories, Burlingame, CA) and confocal images captured using a Leica TCS SP2 confocal system (Leica Microsystems, Wetzlar, Germany).

Total RNA Isolation and RT-PCR

A human podocyte cell line32 was cultured and total RNA prepared as described previously.34 Reverse transcription was done using random hexamers and Superscript III First-Strand Synthesis Supermix (Invitrogen, Carlsbad, CA).32 Sample cDNA (4 μl) was used to perform PCR for APOL1 with the HotStarTaq Master Mix Kit (Qiagen, Valencia, CA) using the following conditions: 95 °C for 15 min; 94 °C for 15 s, 54 °C for 30 s, 72 °C for 30 s (36 cycles); 72 °C for 5 min, 4 °C hold. Primer pairs for APOL1: Forward 5′-CAGCAGTACCATGGACTACGG-3′ and Reverse 5′-CCGCTCCAGTCACAGTTCTTGGTC-3′. For GAPDH message, sample cDNA (1.2 μl) was used as template in the PCR reaction with an annealing temperature of 57 °C and the following primer pair: Forward 5′-GGAGCCAAACGGGTCATC-3′ and Reverse 5′-TGTTGCTGTAGCCGTATTCAT-3′. PCR products were visualized in 1% TBE agarose gel with 0.5 μg/ml ethidium bromide and imaged using a Fotodyne FOTO/analyst archiver workstation (Fotodyne Incorporated, Hartland, WI).

Analysis of APOL1 Protein Expression in Renal Cell Lines and Microvascular Endothelial Cells

Human podocytes, renal proximal tubule cells, and microvascular endothelial cells were grown in culture and whole cell lysates prepared in RIPA buffer (150 mM NaCl, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, 50 mM Tris, pH = 8.0). Endothelial cells were stimulated with either TNF or LPS, or maintained in culture without stimulation as a negative control. Total protein of cell lysates were quantitated using Biorad DC protein assay kit (Bio-Rad Laboratories) according to manufacturer's instructions. Equal quantities of protein lysates were loaded and separated on a 10% SDS-PAGE gel. Proteins were transferred to PVDF membrane blocked in 5% nonfat dry milk in TBS, pH = 7.4, and immunoblotted with anti-APOL1 (Sigma, St. Louis, MO) at 1:500 in 1% nonfat dry milk in TBS, pH = 7.4, with 0.1% tween-20, labeled with HRP-conjugated anti-rabbit secondary antibody, and detected with an enhanced chemiluminescence detection kit (Santa Cruz Biotechnology, Santa Cruz, CA). Membranes were stripped and reprobed with anti-tubulin (Sigma, St. Louis, MO) at 1:1000. Membranes were then stained with Ponceau S. Quantitation of band density was performed using ImageJ software available from the NIH (http://rsbweb.nih.gov/ij/).

Extraction of DNA from Biopsy Material and APOL1 Risk Variant Genotyping

The APOL1 genotypes of renal disease risk variants in the deidentified biopsy samples of individuals with FSGS or HIVAN were determined using DNA extracted from formalin-fixed, paraffin-embedded renal biopsy cores that had been sectioned onto glass slides. A biopsy section from each of the samples used for microscopic analysis was removed from the slide using a sterile scalpel blade, and DNA was extracted using RecoverAll™ Total Nucleic Acid Isolation Kit (Ambion, Inc., Austin, TX) according to manufacturer's instructions. Isolated genomic DNA was quantitated using a Nanodrop-1000 spectrophotometer (Nanodrop Products, Wilmington, DE) and 50–100ng of genomic DNA was used as template in a PCR reaction amplifying a single 380 bp amplicon containing the known risk variants (primers: forward-5′- AGACGAGCCAGAGCCAATC-3′, reverse-5′- CTGCCAGGCATATCTCTCCT-3′). PCR products were evaluated using agarose gel electrophoresis with ethidium bromide; the remainder of the reaction was purified using a PCR purification kit (Marligen Biosciences, Inc., Ijamsville, MD) and sequenced using a 3730xl DNA Analyzer (Applied Biosystems, Carlsbad, CA) with the PCR forward primer. Sequences were evaluated using Sequencher® Software (Gene Codes, Ann Arbor, MI).

DISCLOSURES

None.

Acknowledgments

This work was supported, in whole or in part, by National Institutes of Health grants DK-064719 and DK-071108.

Footnotes

  • Published online ahead of print. Publication date available at www.jasn.org.

  • See related editorial, “Apolipoprotein L1 and the Genetic Basis for Racial Disparity in Chronic Kidney Disease,” on pages 1955–1958.

  • Supplemental information for this article is available online at http://www.jasn.org/.

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APOL1 Localization in Normal Kidney and Nondiabetic Kidney Disease
Sethu M. Madhavan, John F. O'Toole, Martha Konieczkowski, Santhi Ganesan, Leslie A. Bruggeman, John R. Sedor
JASN Nov 2011, 22 (11) 2119-2128; DOI: 10.1681/ASN.2011010069
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