CD44 expression in renal tubular epithelial cells in the kidneys of rats with cyclosporine-induced chronic kidney disease

Abstract

Renal tubular epithelial cell (TEC) injury is the most common cause of drug-induced kidney injury (DIKI). Although TEC regeneration facilitates renal function and structural recovery following DIKI, maladaptive repair of TECs leads to irreversible fibrosis, resulting in chronic kidney disease (CKD). CD44 is specifically expressed in TECs during maladaptive repair in several types of rat CKD models. In this study, we investigated CD44 expression and its role in renal fibrogenesis in a cyclosporine (CyA) rat model of CKD. Seven-week-old male Sprague–Dawley rats fed a low-salt diet were subcutaneously administered CyA (0, 15, or 30 mg/kg) for 28 days. CD44 was expressed in atrophic, dilated, and hypertrophic TECs in the fibrotic lesions of the CyA groups. These TECs were collected by laser microdissection and evaluated by microarray analysis. Gene ontology analysis suggested that these TECs have a mesenchymal phenotype, and pathway analysis identified CD44 as an upstream regulator of fibrosis-related genes, including fibronectin 1 (Fn1). Immunohistochemistry revealed that epithelial and mesenchymal markers of TECs of fibrotic lesions were downregulated and upregulated, respectively, and that these TECs were surrounded by a thickened basement membrane. In situ hybridization revealed an increase in Fn1 mRNA in the cytoplasm of TECs of fibrotic lesions, whereas fibronectin protein was localized in the stroma surrounding these tubules. Enzyme-linked immunosorbent assay revealed increased serum CD44 levels in CyA-treated rats. Collectively, these findings suggest that CD44 contributes to renal fibrosis by inducing fibronectin secretion in TECs exhibiting partial epithelial-mesenchymal transition and highlight the potential of CD44 as a biomarker of renal fibrosis.

Introduction

Drug-induced kidney injury (DIKI) is a major concern in the clinical use of nephrotoxic drugs as well as in the drug development process^{1, 2}. Renal tubular epithelial cells (TECs) are particularly sensitive to drug-induced toxicity because of their pivotal role in the reabsorption and metabolism of xenobiotics^{3, 4}. Acute tubular necrosis is the most common cause of acute kidney injury (AKI)⁵. AKI is associated with a high risk of mortality and is one of the major dose-limiting toxicities in clinical practice^{6, 7}; therefore, toxicological studies on DIKI have mainly focused on AKI. As a result, the identification of TEC injury mechanisms has facilitated the development of AKI-specific biomarkers capable of detecting TEC injuries with high sensitivity and specificity⁸. Moreover, growing epidemiological evidence has demonstrated a relationship between DIKI and chronic kidney disease (CKD)⁹. Therefore, toxicological research on the mechanisms of DIKI chronicity and the development of biomarkers capable of monitoring CKD are urgently required.

TECs have an intrinsic regenerative capacity that contributes to its structural and functional recovery after injury. During the adaptive repair process, dedifferentiated TECs spread to cover the space due to the detachment of necrotic TECs, followed by redifferentiation¹⁰. However, the maladaptive repair of TECs leads to irreversible fibrosis, resulting in CKD^{11, 12, 13}. Studies in the field of toxicologic pathology have shown that the adaptive-repair TECs present a basophilic cuboidal cytoplasm and a high rate of mitosis¹⁴; however, there is limited information regarding the morphology of failed-repair TECs.

In our previous studies, dilated or atrophic tubules were observed in fibrotic lesions in the kidneys of rats with CKD induced by ischemia-reperfusion injury and allopurinol administration^{15, 16} indicating that failed-repair TECs morphologically exhibited dilation and atrophy. In addition, we demonstrated the specific expression of CD44, a transmembrane glycoprotein, in the dilated/atrophic TECs of fibrotic lesions^{15, 16}. CD44 was originally identified as a hyaluronic acid receptor, although subsequent studies have demonstrated that CD44 has multiple functions in several diseases and conditions, including cancer, inflammation, and fibrosis^{17, 18, 19}. Moreover, the degree of CD44 expression and interstitial fibrosis in human kidney diseases, including IgA nephropathy and renal transplantation, is positively correlated^{20, 21}, highlighting the important role of CD44 in renal fibrogenesis. Therefore, further studies are required to evaluate the expression of CD44 and its pathophysiological role in renal fibrogenesis in various animal models of CKD.

In the current study, we examined the morphology of TECs and CD44 expression in the kidneys of rats with CKD induced by cyclosporine (CyA), an immunosuppressant with nephrotoxicity in humans and experimental animals^{22, 23}. To clarify the pathophysiology of CD44, TECs of fibrotic lesions were collected by laser microdissection (LMD) and subjected to microarray analysis. Additionally, serum CD44 levels were measured to investigate the potential of CD44 as a biomarker for renal fibrosis.

Materials and Methods

Chemicals

CyA (purity: ≥97%) and olive oil were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan) and Wako Pure Chemical Industries (Tokyo, Japan), respectively.

Experimental animals and housing conditions

Specific pathogen-free Sprague–Dawley (Crl:CD(SD)) rats (males, 5-weeks-old) were obtained from The Jackson Laboratory Japan, Inc. (Kanagawa, Japan) and acclimated for 1 week prior to experimentation. The animals were housed in polycarbonate cages with soft chip bedding (Sankyo Labo Service, Tokyo, Japan) in a room with a barrier system and a controlled light/dark cycle (12 h), ventilation (air-exchange rate: 20 times/h), temperature (23 ± 1 °C), and relative humidity (50% ± 5%) during the study. All animals had free access to food and water. Animal experiments were approved by the Animal Care and Utilization Committee of the National Institute of Health Sciences (protocol no. 848).

Animal treatment

Fifteen rats were fed a low-salt diet (0.05% sodium; CLEA Japan, Inc., Tokyo, Japan) throughout the experimental period, as described previously²². After 1 week, rats were subcutaneously administered olive oil (vehicle, 4 mL/kg) or 15 or 30 mg/kg CyA (4 mL/kg) once daily for 4 weeks (n=5/group). The CyA dose was determined based on a previous study²². On the day after the final administration, rats were euthanized by exsanguination via transection of the abdominal aorta under deep anesthesia induced by isoflurane inhalation. At necropsy, blood samples were collected from the abdominal aorta to evaluate blood urea nitrogen and serum creatinine levels using serum biochemistry. Serum CD44 levels were measured using a rat CD44 enzyme-linked immunosorbent assay (ELISA) kit (Novus Biologicals, Centennial, CO, USA), according to the manufacturer’s instructions. The kidneys were sampled and weighed. Kidney tissues were cut into pieces, fixed in 10% neutral buffered formalin at room temperature overnight, embedded in paraffin, and sectioned (4-μm thickness) for hematoxylin and eosin (HE) and Sirius red staining. Additional portions of kidney tissues were embedded in O.C.T. compound (Sakura Fintech Japan, Tokyo, Japan) and stored at –80°C for LMD and microarray analysis. Residual pieces of kidney tissues were immediately frozen in liquid nitrogen and stored –80°C for reverse transcription quantitative polymerase chain reaction (RT-qPCR) and western blotting analysis.

Immunohistochemistry (IHC)

Formalin-fixed paraffin-embedded sections were autoclaved at 121°C for 15 min in 10 mM citrate buffer (pH 6.0) for antigen retrieval. The sections were then incubated in 3% H₂O₂/methanol to inactivate endogenous peroxidase activity, followed by incubation with 10% normal goat serum at room temperature for 30 min. Subsequently, the sections were incubated with antibodies against CD44 (1:1,000, rabbit polyclonal; Abcam, Cambridge, UK), Ki67 (1:500, rabbit polyclonal; Abcam), α-smooth muscle actin (αSMA; 1:50, Agilent Technologies, Santa Clara, CA, USA; clone: 1A4), aquaporin-1 (AQP1; 1:300; Santa Cruz, Dallas, TX, USA; clone: B-11), vimentin (1:200; Agilent Technologies; clone: V9), collagen type IV (1:400, rabbit polyclonal; Abcam), and fibronectin (1:800, rabbit polyclonal; Abcam) at 4°C overnight. Following this, secondary antibody reactions were performed using a horseradish peroxidase (HRP)-conjugated antibody (Histofine Simple Stain MAX PO; Nichirei Bioscience Inc., Tokyo, Japan). 3,3′-Diaminobenzidine tetrahydrochloride was applied for visualization.

Image analysis

Specimens for Sirius red staining and IHC were converted to whole-slide images using Aperio AT2 (Leica Biosystems, Wetzlar, Germany) and captured using HALO image analysis software (Indica Labs, Albuquerque, NM, USA). All tubules positive for CD44 and vimentin in the cortex and outer stripe of the outer medulla (OSOM) in the whole kidney tissue were enumerated, and the number of tubules positive for AQP1 was counted in 10 randomly selected fields (five areas each in the cortex and OSOM) using a 20× objective lens. The number of tubules positive for CD44, vimentin, and AQP1 per unit area was calculated. The tissue classifier module was used to quantify the percentage of positive areas for Sirius red and IHC staining of fibronectin.

Immunofluorescence (IF)

To analyze the co-localization of CD44 and other factors, double IF staining with two antibodies derived from different hosts (AQP1 and vimentin) or the same host (fibronectin and collagen type IV) was performed as previously reported¹⁶.

Formalin-fixed, paraffin-embedded sections (4-μm-thick) were autoclaved at 121°C for 15 min with 10 mM citrate buffer (pH 6.0) for antigen retrieval. For double staining of AQP1 and vimentin, blocking was performed with 10% normal donkey serum and the sections were incubated with either mouse anti-AQP1 antibody (1:300) or mouse anti-vimentin antibody (1:200) at 4°C overnight. Sections were then incubated with secondary anti-mouse immunoglobulin (Ig) G antibodies (1:200; Alexa Fluor 647; Abcam) at room temperature for 2 h. Subsequently, the sections were incubated with rabbit antibodies against CD44 (1:500) at 37°C for 1 h, followed by incubation with secondary anti-rabbit IgG antibodies (1:1,000; Alexa Fluor 488; Abcam).

Double staining of fibronectin and collagen type IV was performed using the tyramide signal amplification (TSA) method. In brief, after antigen retrieval, the sections were immersed in 3% H₂O₂/methanol to inactivate endogenous peroxidase activity. Blocking was performed using 10% normal goat serum, and sections were incubated with rabbit antibodies against CD44 (1:4,000) at 4°C overnight. Secondary antibody reactions were performed using a HRP-conjugated antibody (Histofine Simple Stain MAX PO; Nichirei Bioscience, Inc.). Sections were incubated with TSA Plus working solution from the TSA Plus Fluorescein Kit (PerkinElmer, Waltham, MA, USA) for HRP-catalyzed deposition of fluorophore-labeled tyramide in kidney tissues. To strip anti-CD44 antibodies and secondary antibodies from the tissues while retaining fluorophore-labeled tyramide deposition, sections were autoclaved at 121°C for 15 min with 10 mM citrate buffer (pH 6.0). After blocking with 10% normal donkey serum, sections were incubated at 37°C for 1 h with rabbit antibodies against fibronectin (1:400) or collagen type IV (1:200), followed by incubation with secondary anti-rabbit IgG antibodies (1:1,000; Alexa Fluor 647; Abcam).

A Vector TrueVIEW Autofluorescence Quenching Kit (Vector Laboratories, Burlingame, CA, USA) was applied to all IF sections to eliminate autofluorescence, and the sections were mounted with Vibrance Antifade Mounting Medium containing DAPI (Vector Laboratories). The IF images were obtained using an All-in-One Fluorescence Microscope BZ-X710 (Keyence Corp., Osaka, Japan).

In situ hybridization (ISH)

Formalin-fixed, paraffin-embedded sections (4-μm-thick) were used for ISH. Localization of fibronectin 1 (Fn1) mRNA was detected using a specific probe (probe number: 520411) from an RNAscope 2.5 HD Reagent Kit Brown (Advanced Cell Diagnostics, Newark, NJ, USA) according to the manufacturer’s instructions.

LMD for sampling of TECs and tRNA extraction

Sampling of TECs by LMD and extraction of tRNA were performed as previously reported^{16, 24}. Briefly, O.C.T.-embedded frozen kidney tissues were cut into 16-µm-thick sections using a cryotome (Leica Microsystems, Wetzlar, Germany), fixed with 5% acetic acid/ethanol, and subjected to HE staining. An LMD6000 system (Leica Microsystems) was used to capture images of the normal proximal renal tubules in the control group and the atrophic, dilated, or hypertrophic renal tubules in fibrotic lesions in the 30 mg/kg CyA group (n=4/group). Total RNA from normal proximal renal tubules and renal tubules in fibrotic lesions (a mixture of atrophic, dilated, and hypertrophic renal tubules) was extracted using an RNeasy Micro Kit (Qiagen GmbH, Hilden, Germany). The concentration and quality of total RNA were analyzed using an Agilent 2100 Bioanalyzer (Agilent Technologies). RNA integrity number (RIN) was used as an index of RNA degradation. RNA samples with RIN value >7.0 was amplified and synthesized into SPIA cDNA using an Ovation PicoSL WTA System V2 (NuGEN, San Carlos, CO, USA) for microarray analysis.

Microarray and data analysis

Microarray analysis was performed as previously reported^{16, 24}. Briefly, SPIA cDNA was labeled with cyanine-3 dye using the SureTag DNA Labeling Kit (Agilent Technologies). cDNA concentration, dye incorporation, and quality were analyzed using an ultraviolet-visible spectrophotometer. Fluorescent-labeled cDNA was hybridized to Agilent 4 × 44 K Whole Rat Genome Microarray gene expression chips (version 3.0; Agilent Technologies) following the manufacturer’s protocol. Hybridized microarray chips were scanned using an Agilent Microarray Scanner (Model G256BA; Agilent Technologies). Feature Extraction software (Agilent Technologies) was used for image analysis and data extraction. Normalization of gene expression data and filtering of probe sets by expression levels, flags, and errors were performed using the GeneSpring software (Agilent Technologies). Differences in gene expression between normal TECs and TECs in fibrotic lesions were determined by statistical analysis (analysis of variance, cutoff value: p<0.05; t-test: Benjamini–Hochberg false discovery rate) and fold-change (>2.0 or <–2.0). Gene ontology (GO) analysis (cutoff value: p<0.05) and Ingenuity Pathway Analysis (IPA; Qiagen) were performed on the extracted genes.

Quantitative reverse transcription polymerase chain reaction (RT-qPCR)

Total RNA was extracted from freshly frozen kidney tissues using an RNeasy Mini Plus kit (Qiagen). To analyze the mRNA expression of total Cd44, including all splice variant isoforms of CD44, RT-qPCR assays for rat total Cd44 (Rn00681157_m1; Applied Biosystems, Waltham, MA, USA) targeting invariable exons 16–17 were performed using the following primers: forward primer, 5′-CCCTGGCCACCAGTGATG-3′ (exon boundary 5–16); reverse primer, 5′-TCCCACTTGAGTGTCCAGCTAA-3′ (exon boundary 16–17). Analysis of the CD44v3-10 (CD44v3) expression was performed using the following primers: forward primer, 5′-GGCCACCAGTACGGAGTCAA-3′ (exon boundary 5–8); reverse primer, 5′-TTGCAATGGTGCTGGAGATAAA-3′ (exon boundary 8–9). Primers for eukaryotic 18S rRNA (Hs99999901_s1; Agilent Technologies) were used as endogenous references. PCR was performed using a QuantStudio 3 Real-Time PCR System (Applied Biosystems) with TaqMan Fast Universal PCR Master Mix and TaqMan Gene Expression Assays (Life Technologies, Carlsbad, CA, USA). Expression levels of the target genes were calculated according to the relative standard curve method and normalized to 18S rRNA expression. Data are presented as fold-change values obtained by comparing the expression of genes in the kidneys of CyA-treated rats to those in the control group.

Western blotting analysis

Frozen kidneys were homogenized using radioimmunoprecipitation assay buffer, and the resulting supernatants were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred to 0.45-µm polyvinylidene difluoride membranes (Millipore, Billerica, MA, USA). To block non-specific reactions, the membranes were incubated with 5% bovine serum albumin for 1 h at room temperature. Rabbit polyclonal antibodies against CD44 (1:1,000 dilution; Abcam) were used to detect the target protein, and rabbit polyclonal antibodies against β-actin (1:1,000 dilution; Cell Signaling Technology, Danvers, MA, USA) were used as an endogenous control for kidney samples. Membranes were incubated with primary antibodies overnight at 4°C, followed by incubation with secondary antibodies (Cell Signaling Technology) for 60 min at room temperature. Protein detection was facilitated by chemiluminescence using ECL Prime (GE Healthcare, Tokyo, Japan).

Statistical analysis

Statistical analyses were performed using GraphPad Prism version 9 (GraphPad Software Inc., San Diego, CA, USA). The results are presented as means ± standard deviations (SDs). The significance of differences was analyzed using analysis of variance, followed by Dunnett’s multiple comparison test for comparisons of data. Correlations between two parameters were analyzed using Spearman’s correlation test. P<0.05 was considered statistically significant.

Results

Histopathological analysis

Kidney weights and expression levels of serum biomarkers for kidney injury were significantly higher in the CyA groups than in the control group (Supplementary Table 1). The Sirius red-positive area was significantly increased in the 15 and 30 mg/kg CyA groups compared with that in the control group (Fig. 1A), and the number of αSMA-positive myofibroblasts was increased in the interstitium of fibrotic lesions in CyA groups (Supplementary Fig. 1A). In the fibrotic lesions of the CyA groups, renal tubules were atrophied, dilated, or hypertrophied, and expressed CD44 in the cytoplasmic membrane of their TECs (Fig. 1B). These tubules also had basophilic cytoplasm and were surrounded by a slightly thickened basement membrane. The increase in number of Ki67-positve nuclei in these tubules were not increased compared to normal tubules of the control group (Supplementary Fig. 1B). Furthermore, atrophied, dilated, or hypertrophic tubules were mainly located in the fibrotic lesions and were barely detected in the non-fibrotic areas. Fibrotic lesions with these tubules were widely observed in the cortex and OSOM. The number of CD44-positive tubules significantly increased in both CyA groups and positively correlated with the Sirius red-positive area (Fig. 1C). CD44 was also slightly expressed in parietal epithelial cells of the control and CyA groups. In addition, vacuolation, and necrosis of TECs and calcification were observed in the CyA groups (Supplementary Fig. 1C). Inflammatory cell infiltration into the interstitium was also detected in the CyA group, and some inflammatory cells were positive for CD44.

Fig. 1.

Morphology and CD44 expression of renal tubules in fibrotic lesions. (A) Representative photographs of Sirius red staining and image analysis of the Sirius red-positive area. (B) Hematoxylin and eosin (HE) staining and immunohistochemistry for CD44. (C) The number of CD44-positive tubules and correlation with the Sirius red-positive area. Values are presented as mean ± SDs for 5 rats (A and C). Correlation plots include the data for 15 rats (C). **: Significantly different from the control group at p<0.01. ^#: Significant correlation between two parameters at p<0.05.

Microarray analysis

Figure 2A shows representative photographs for sampling of normal tubules in the control group and atrophic, dilated, and hypertrophic renal tubules in fibrotic lesions from the 30 mg/kg CyA group using LMD. Microarray analysis revealed 930 upregulated genes and 1,161 downregulated genes in TECs of fibrotic lesions in the 30 mg/kg CyA group compared to normal TECs in the control group.

Fig. 2.

Microarray analysis of renal tubules in fibrotic lesions. (A) Representative photographs for sampling of normal renal tubules in the control group and atrophic, dilated, and hypertrophic renal tubules in the 30 mg/kg CyA group using laser microdissection (LMD). Arrowheads: renal tubules subjected to sampling by LMD. (B, C) The results of Gene ontology analysis (B) and Ingenuity Pathway Analysis (C). **: Corrected p<0.01.

GO terms in the upregulated and downregulated gene sets in TECs of fibrotic lesions were sorted according to corrected p-values (Supplementary Tables 2 and 3). A total of 199 and 378 GO terms were present for the upregulated and downregulated genes, respectively. Among these GO terms, we focused on those related to the extracellular matrix (ECM), transporters, and metabolism. The top 4 ECM-related GO terms in the up-regulated gene set and the top 4 GO terms related to transporters and metabolism in the down-regulated gene set were extracted. The numbers of upregulated and downregulated genes included in each GO term were counted. The results showed an increase in the number of upregulated genes related to the ECM, but a decrease in the number of downregulated genes related to transporters and metabolism in TECs of fibrotic lesions (Fig. 2B).

In IPA, upstream regulator analysis was performed to identify the genes downstream of CD44, and the biological significance of the extracted downstream genes was investigated using disease and function analyses. Several fibrosis-associated genes downstream of CD44, including Fn1, were identified (Fig. 2C).

Analysis of epithelial-mesenchymal transition (EMT)-related factors

AQP1, a differentiation marker of proximal renal tubules, was abundantly expressed in the tubules of the control group, whereas its expression was decreased or disappeared in tubules of fibrotic lesions (Fig. 3A). The number of AQP1-positive tubules was significantly decreased in the CyA group and negatively correlated with the number of CD44-positive tubules (Fig. 3B). IF analysis demonstrated that the expression of AQP1 and CD44 was mutually exclusive in the CyA groups (Fig. 3C). The expression of vimentin, a mesenchymal marker, was barely detectable in normal renal tubules and was clearly increased in renal tubules of fibrotic areas (Fig. 3D). The number of vimentin-positive tubules was significantly increased in the CyA groups and positively correlated with the number of CD44-positive tubules (Fig. 3E). IF revealed the co-expression of vimentin and CD44 in the renal tubules of the CyA groups (Fig. 3F). IHC for collagen type IV showed a thin basement membrane surrounding the normal renal tubules of the control group, whereas CD44-positive renal tubules of the CyA groups were surrounded by a slightly thickened basement membrane (Fig. 3G and 3H).

Fig. 3.

Immunohistochemistry (IHC) for differentiation and dedifferentiation markers of renal tubules and the basement membrane. (A) Representative photographs of IHC for aquaporin-1 (AQP1). (B) The number of AQP1-positive tubules and correlation with the number of CD44-positive tubules. (C) Representative photographs of immunofluorescence (IF) for AQP1 and CD44. (D) Representative photographs of IHC for vimentin. (E) The number of vimentin-positive tubules and correlation with the number of CD44-positive tubules. (F) Representative photographs of IF for vimentin and CD44. (G, H) Representative photographs of IHC for collagen type IV (Col IV, G) and IF for Col IV and CD44 (H). Values are presented as mean ± SDs for 5 rats (B and E). Correlation plots include the data for 15 rats (B and E). *, **: Significantly different from the control group at p<0.05 and p<0.01, respectively. ^##: Significant correlation between two parameters at p<0.01.

Localization of Fn1 mRNA and fibronectin protein

The results of ISH of Fn1 mRNA and IHC of fibronectin are shown in Fig. 4A. In the control group, Fn1 mRNA was expressed in stromal cells and was barely detectable in normal renal tubules, whereas fibronectin protein was weakly expressed in the interstitium. In contrast, Fn1 mRNA expression increased in the cytoplasm of renal tubules in fibrotic lesions of the CyA group, and fibronectin protein expression was increased in the interstitium. Stromal cells in the fibrotic lesions of the CyA group also showed an obvious increase in Fn1 mRNA expression. The fibronectin-positive area was significantly increased in the CyA groups and positively correlated with the number of CD44-positive tubules (Fig. 4B). IF revealed increased fibronectin expression in the interstitium around the CD44-positive renal tubules (Fig. 4C).

Fig. 4.

Localization of fibronectin 1 (Fn1) mRNA and fibronectin protein. (A) Representative photographs of in situ hybridization for Fn1 and immunohistochemistry for fibronectin. (B) Image analysis of the fibronectin-positive area and correlation with the number of CD44-positive tubules. (C) Representative photographs of immunofluorescence for fibronectin and CD44. Values are presented as mean ± SDs for 5 rats (B). Correlation plots include the data for 15 rats (B). **: Significantly different from the control group at p<0.01. ^##: Significant correlation between two parameters at p<0.01.

Evaluation of CD44 levels in serum

ELISA demonstrated an increasing trend and a significant increase in serum CD44 levels in the 15 and 30 mg/kg CyA groups, respectively (Fig. 5A). CD44 levels also positively correlated with the number of CD44-positive tubules or Sirius red-positive areas in the kidneys (Fig. 5B).

Fig. 5.

Expression of CD44 in the serum. (A) CD44 levels in serum. (B) Correlation between CD44 levels in the serum and the number of CD44-positive tubules or the Sirius red-positive area in the kidneys. Values are presented as mean ± SDs for 5 rats (A). Correlation plots include the data for 15 rats (B). *: Significantly different from the control group at p<0.05. ^#, ^##: Significant correlation between two parameters at p<0.05 and p<0.01, respectively.

Splice variant analysis of CD44 expressed in the kidneys

RT-qPCR results are shown in Fig. 6A. The mRNA expression of total Cd44 and CD44standard were significantly increased to the same degree in the CyA groups. The expression of CD44v3 was also significantly increased in the CyA group, but the levels were lower than those of total Cd44 and CD44standard. Western blotting showed that CD44-specific bands were clearly detected at approximately 82 kDa in the CyA groups (Fig. 6B).

Fig. 6.

Analysis of the expression of alternatively spliced variants of CD44. (A) RT-qPCR analysis of the expressions of total CD44, CD44standard, and CD44v3. (B) Western blotting analysis of CD44 expression in the kidneys. Values are presented as mean ± SDs for 5 rats (A). **: Significantly different from the control group at p<0.01.

Discussion

Failed-repair TECs have been typically considered passive victims or bystanders during CKD development; however, recent evidence has clearly demonstrated their active role in renal fibrogenesis^{25, 26}. In a previous study, we showed that CD44 was expressed in failed-repair TECs in rat CKD models induced by ischemia-reperfusion injury and allopurinol administration^{15, 16}. Because several reports have demonstrated multiple functions of CD44 in both physiological and pathological conditions²⁷, we investigated the significance of CD44 expression in the pathophysiology of renal fibrosis using a CKD animal model.

CyA, a calcineurin inhibitor, has been widely used as an immunosuppressant in patients with autoimmune diseases and in those who have undergone organ transplantation. However, CyA causes acute and chronic kidney injury²⁸. In previous reports, researchers developed rat models of CKD using a combination of CyA administration and a low-salt diet, in which activation of the renin-angiotensin system is involved in renal toxicity and fibrosis^{22, 29}. In the current study, CyA administration induced the formation of lesions exhibiting TEC vacuolation, calcification, and fibrosis in the kidneys of rats fed a low-salt diet, which is consistent with the results of a previous report³⁰. Detailed histopathological examination also revealed atrophic, dilated, and hypertrophic TECs in fibrotic lesions. Since the abnormal repair of TECs leads to renal fibrosis¹¹, the TECs observed in fibrotic lesions in the current study were thought to be involved in the process of maladaptation. In our previous studies using rat models of CKD induced by ischemia-reperfusion injury and allopurinol administration, dilated and atrophic TECs were observed in fibrotic lesions, whereas hypertrophic TECs were not detected^{15, 16}. Further analyses are required to clarify the mechanisms underlying the development of TECs with failed repair.

IHC demonstrated that CD44 was expressed in the atrophic, dilated, and hypertrophic TECs of fibrotic lesions. We also demonstrated that regenerative tubules were negative for CD44 in the kidneys of rats subjected to ischemia reperfusion injury¹⁵. These observations supported the specific expression of CD44 in failed-repair TECs. Since fibrotic lesions with CD44-positive tubules were widely distributed in the cortex and OSOM, which are largely composed of proximal renal tubules, and their number was negatively correlated with that of AQP1-positive renal tubules, a large portion of the atrophic, dilated, or hypertrophic tubules in fibrotic lesions could have originated from the proximal renal tubules. To further investigate the characteristics of TECs expressing CD44, TECs in fibrotic lesions were collected using LMD and subjected to microarray analysis. The results of GO analysis suggested that TECs in the fibrotic lesions showed a loss of epithelial characteristics and acquisition of a mesenchymal phenotype, indicating that EMT occurred in these TECs. TECs exhibiting EMT have long been considered to migrate into the interstitium and transform into myofibroblasts that produce ECM^{31, 32, 33}. However, recent lineage-tracing studies have revealed a minor contribution of TECs to the pool of myofibroblasts^{34, 35, 36}. Additionally, TECs acquire a mesenchymal phenotype during attachment to the basement membrane^{37, 38, 39}, which has been proposed as a new concept termed partial epithelial-mesenchymal transition (pEMT)^{40, 41}. TECs exhibiting pEMT activate fibroblasts via secretion of profibrotic cytokines or growth factors⁴¹. In the current study, IHC revealed that TECs in fibrotic lesions showed decreased expression or absence of differentiation markers, whereas mesenchymal markers were clearly expressed. IF results demonstrated the mutually exclusive expression of CD44 and AQP1, as well as the co-expression of CD44 and vimentin. In contrast, TECs in fibrotic lesions were surrounded by the basement membrane, and there was no evidence suggesting migration of CD44-positive TECs into the interstitium. These results suggest that pEMT could have occurred in CD44-positive TECs during the development of fibrosis in this CKD model.

Pathway analysis using microarray data suggested that CD44 may be an upstream factor for fibrosis-related genes, including Fn1, which encodes fibronectin. Taken together with the results of GO analysis, these findings suggest that CD44-positive TECs produce ECM. To confirm this hypothesis, we analyzed Fn1 mRNA expression using ISH, which is a useful method for detect secretory cells^{42, 43, 44}. The results showed that Fn1 expression was increased in the cytoplasm of TECs in fibrotic lesions. In addition, IHC detected fibronectin protein in the interstitium around these TECs. Overall, these results suggested that CD44 induces ECM production in TECs exhibiting pEMT, which may contribute to renal fibrosis. Numerous in vitro studies have demonstrated that ECM is produced by TECs^{45, 46, 47}, but there is no evidence establishing ECM secretion by TECs in vivo. Thus, further investigations are required to confirm ECM production by CD44-positive TECs. In the present study, ISH clearly demonstrated an increase in Fn1 mRNA expression in stromal cells, presumably myofibroblasts. Since myofibroblasts contribute mainly to renal fibrogenesis via the production of various types of ECM components, such as fibronectin, collagen type I, and collagen type III³³, the contribution of fibronectin from TECs exhibiting pEMT to the progression of renal fibrosis needs to be further examined. In a previous study using a rat model of CKD induced by allopurinol, we demonstrated similar results, indicating that CD44 induces ECM production in TECs expressing pEMT¹⁶. Therefore, the pathophysiological role of CD44 in renal fibrogenesis is common, regardless of the cause of primary TEC injury.

ELISA revealed increased serum CD44 levels in the CyA groups. In addition, serum CD44 levels positively correlated with the number of CD44-positive tubules in the kidney, indicating that the increase in serum CD44 levels correlated with the induction of CD44 expression in the TECs of fibrotic lesions. Thus, renal fibrosis could be monitored by measuring serum CD44 levels. We have also demonstrated the expression of CD44 in TECs immediately after AKI, prior to fibrosis¹⁵. Since typical kidney injury biomarkers monitoring renal function transiently decrease after AKI in the process of CKD progression^{48, 49, 50}, we are currently investigating CD44 expression in the serum immediately after AKI to evaluate the potential of CD44 as a biomarker for predicting the chronicity of AKI. Several reports have demonstrated the detection of CD44 in the serum of patients with various types of tumors and liver disease^{18, 51, 52, 53, 54}. CD44 is a single-pass transmembrane glycoprotein consisting of an extracellular, a transmembrane, and an intracellular domain⁵⁵. The extracellular domain of CD44 can be cleaved by metalloproteinases such as membrane-type 1 matrix metalloproteinase and ADAM10 and is closely related to the proliferation and migration of cancer cells^{56, 57}. The soluble fragment of the cleaved extracellular domain of CD44 can be detected in the serum and other bodily fluids⁵⁸. Notably, Masson et al. demonstrated that the full-length CD44 standard isoform is present in the serum of patients with colorectal cancer⁵². Thus, multiple mechanisms may be involved in CD44 degradation and leakage into the bodily fluids. In contrast to the results of our previous study using a rat model of CKD induced by allopurinol¹⁶, CD44 was not detected in the urine of rats administered CyA by western blotting analysis in the current study (data not shown). Although the sensitivity of CD44 detection by western blotting may explain this discrepancy, further investigation is required to verify the potential of CD44 as a urinary biomarker for CKD.

CD44 has several alternately spliced variants. Rodent Cd44 consists of 10 invariable exons (exons 1–5 and 16–20) and 10 variable exons (exons 6–15)⁵⁹. CD44standard is the shortest isoform with no variable exons, whereas CD44 variant isoforms contain any combination of variable exons with 10 invariable exons. The variable domain, located in the extracellular region, alters the functional role of CD44 variant isoforms by promoting their interaction with several types of microenvironmental factors^{59, 60}. In the current study, qPCR and western blot analysis suggested that the main isoform of CD44 expressed in the kidneys was the CD44standard, with a molecular weight of approximately 82 kDa. Rampanelli et al. showed that overexpression of CD44standard or CD44v3 in systemic organs leads to exacerbation or attenuation of renal fibrosis, respectively, in a mouse model of renal fibrosis induced by unilateral ureteral obstruction⁶¹.

In conclusion, our findings suggest that CD44 induces ECM production and secretion in TECs exhibiting pEMT, which may contribute to renal fibrogenesis. In addition, because serum CD44 levels increased in rats treated with CyA, CD44 could be a biomarker capable of detecting renal fibrosis.

Disclosure of Potential Conflicts of Interest

All authors declare that they have no conflicts of interest.