odd skipped-related 2 as a novel mark for labeling the proximal convoluted tubule within the zebrafish kidney

Abstract

The proximal convoluted tubule (PCT) of the kidney is a crucial functional segment responsible for reabsorption, secretion, and the maintenance of electrolyte and water balance within the renal tubule. However, there is a lack of a well-defined endogenous transgenic line for studying PCT morphogenesis. By analyzing single-cell transcriptome data from the adult zebrafish kidney, we have identified the expression of odd-skipped-related 2 (osr2, which encodes an odd-skipped zinc-finger transcription factor) in the PCT. To gain insight into the role of osr2 in PCT morphogenesis, we have generated a transgenic zebrafish line Tg(osr2:EGFP), expressing enhanced green fluorescent protein (EGFP). The EGFP expression pattern closely mirrors that of endogenous Osr2, faithfully recapitulating its native expression profile. During kidney development, we can use EGFP to track PCT development, which is also preserved in adult zebrafish. Additionally, osr2:EGFP-labeled zebrafish PCT fragments displayed short lengths with infrequent overlap, rendering them conducive for nephrons counting. The generation of Tg(osr2:EGFP) transgenic line is accompanied by simultaneous disruption of osr2 activity. Importantly, our findings demonstrate that osr2 inactivation had no discernible impact on the development and regeneration of Tg(osr2:EGFP) zebrafish nephrons. Overall, the establishment of this transgenic zebrafish line offers a valuable tool for both genetic and chemical analysis of PCT.

1. Introduction

The PT (proximal tubule) is a critical component of the nephron and performs the majority of reabsorption and processing of the glomerular filtrate. It consists of two distinct segments: PCT and the proximal straight tubule (PST). In mammals, the PCT is characterized by its convoluted structure, while the PST is straight and lacks convolutions. Functionally, both segments share similar roles, including the reabsorption of water, glucose, amino acids, and ions, as well as the excretion of waste products [[1], [2], [3]]. However, these functions are mainly contributed by PCT, while the PST's contribution is relatively limited [4]. Injured PCT cells play a pivotal role in the pathogenesis of acute kidney injury (AKI), chronic kidney disease, and diabetic kidney disease [5,6]. Considering the high conservation of genes between zebrafish and mammalian PCT [7], zebrafish models hold great value for the study of PCT development and diseases.

Tissue-specific transgenic technology has emerged as an invaluable tool for studying tissue development. In zebrafish, several transgenic lines with fluorescent protein have been constructed to visualize kidney organ morphogenesis. For instance, the transgenic line Tg(cdh17:DsRed) (the line ID: ZDB-FIG-230223-3) labels renal tubules [8]. The Tg(dβh:GFP) zebrafish line utilizes the dopamine-β-hydroxylase (dβh) promoter to drive GFP expression, marking PT development [9]. Another transgenic line, Tg(ret1:GFP), exhibits GFP expression driven by the ret loci, marking the most distal pronephric duct [10]. The ET33-D10 zebrafish line with GFP expression marks the PT [11]. Additionally, the Tg(gtshβ:GFP) zebrafish line, featuring green fluorescent protein under the control of the grouper gtshβ promoter, provides the details of PT development within the transgenic zebrafish line [12]. Despite these advances, specific endogenous transgenic lines for visualizing PCT development and morphogenesis are still lacking.

The osr2 gene, which encodes a zinc-finger protein, is detected in higher vertebrates such as mouse [13]. Studies on mouse osr2 mutant lines have revealed their essential role in proper patterning of the endoderm, palate, bones, teeth, and synovial joints in the limbs [[14], [15], [16], [17]]. Notably, osr2 has been identified as a key regulator of palate development and a potential candidate gene associated with human cleft palate formation [18,19]. Furthermore, osr2 is expressed in the kidney, and its presence has been observed in the genomes of Xenopus and zebrafish, where it appears to contribute, to some extent, to kidney formation [20,21]. A previous study demonstrated the requirement of osr2 for zebrafish pronephros development, as evidenced by the expression of osr2 in the pronephric kidney and the occurrence of pericardial edema in osr2 knockdown embryos [20]. However, conflicting reports indicate that osr2 knockdown embryos do not exhibit abnormalities in kidney development [22]. Therefore, further evidence is needed for the specific effects of osr2.

In this study, we successfully generated a transgenic zebrafish line, Tg(osr2:EGFP), utilizing the CRISPR/Cas9 knock-in technology. Our findings demonstrate that osr2 serves as a reliable marker for PCT. The EGFP expression in the Tg(osr2:EGFP) kidney enables labeling of PCT. Importantly, the complexity of nephrons poses challenges in their counting. Traditionally, in situ hybridization has been employed for nephron counting; however, this method is time-consuming and intricate. In contrast, the osr2:EGFP-labeled fragments displayed short lengths and infrequent overlap, providing a practical advantage for efficient nephron counting. Furthermore, our investigations revealed that osr2 inactivation in Tg(osr2:EGFP) transgenic fish did not adversely affect the development and regeneration of the kidneys, showing results comparable to the control group. Overall, our study highlights the utility of the Tg(osr2:EGFP) zebrafish line as an innovative resource for studying PCT development, regeneration, and related research in a genetic or chemical context.

2. Materials and methods

2.1. Zebrafish maintenance

Adult and embryonic zebrafish were reared following established protocols outlined in the zebrafish book [23]. The adult zebrafish selected for experiments ranged in age from 3 to 12 months. These zebrafish were maintained under standard conditions, subjected to a 10-h dark cycle and a 14-h light cycle, and were fed twice daily. Before performing experiments, zebrafish individuals were anesthetized using 0.0168% buffered tricaine (MS-222, Sigma). The AB strain (ZFIN ID: ZDB-GENO-960809-7) of zebrafish was employed as the wild-type (WT) for line generation. For this study, we utilized transgenic zebrafish lines expressing the GFF (Gal4FF, Gal4 transcription activator) and the EGFP [24,25]. This transgenic line was generated through CRISPR/Cas9 knock-in technology, as previously described [24,25]. Additionally, the cdh17:DsRed zebrafish line was established in our previous study [8]. The animal care and use protocol employed in this study received approval from the Institutional Animal Care and Use Committee of the Army Medical University, China (SYXK-PLA-2007035).

2.2. Single-cell gene expression pattern analysis

To analyze the expression pattern of osr2 within zebrafish kidneys, we employed a previously published single-cell RNA-sequencing dataset (GSE100910) [26]. For the purpose of data preprocessing, normalization, dimensionality reduction, and subsequent clustering analysis (resolution = 0.4), we applied the Seurat package (version 4.2.0), a widely utilized computational tool in the field of single-cell genomics. PTEC was further classified into PCT and PST by R packages, including ggplot2 (version 3.3.6), dplyr (version 2.2.1), Seurat (version 4.2.0), and tidyverse (version 1.3.2) for dimensionality reduction, and subsequent clustering analysis (resolution = 0.2). This classification was based on marker genes that are specific and highly expressed in PCT (slc20a1a) and PST (slc13a1 and slc13a3) [27].

2.3. Production of sgRNA and zCas9 mRNA

The zCas9 expression plasmid, pGH-SP6-zCas9 [1], was enzymatically linearized using XbaI and subsequently utilized for in vitro synthesis of Cas9 mRNA employing the mMACHINE SP6 Ultra kit (Ambion). The resulting zCas9 mRNA was purified utilizing the same kit. According to previously described criteria [2], the single guide RNA (sgRNA) was designed. To minimize potential off-target effects, the CRISPR/Cas9 design tool (http://chopchop.cbu.uib.no/) was employed to identify specific target sequences. The sequences of the designed sgRNA is 5′-TCACACTTTTACCCCGGGGTTGG-3’. The sgRNA was annealed and subsequently cloned into the PSP6-sgRNA vector downstream of the SP6 promoter. To synthesize the sgRNA, the MAXIscript SP6 Kit (Ambion) was utilized, and purification of the sgRNA was performed using the mirVana™ miRNA Isolation Kit (Ambion).

2.4. Cloning of donor plasmids

The donor plasmid consists of four parts: a right arm, P2A-GFF, a left arm, and cryaa-EGFP. The right and left arms corresponding to osr2 were amplified by the KOD-PLUS Neo DNA polymerase from genomic DNA of WT zebrafish (right arm Forward: 5′-ATGCGGTGGGCTCTATGGAAACCCTTCAAGTGTCAGGAGTG-3’; right arm Reverse: 5′-GAATAGGAACTTCGGTACGGCGGCGGCATGTGTTTGTGT-3’; 840 bp; left arm Forward: 5′-GTAAAACGACGGCCAGTGGCAGACTCCCGGCCAAGAGC-3’; left arm Reverse: 5′-ATTAGTAGCTCCGGATCCCTCCTTAGAGTGGATATATCTGAGGAAC-3’; 840 bp). Subsequently, the amplified arms were individually inserted into the 3′ and 5′ regions of the P2A-GFF fragment, leading to the generation of the donor plasmid.

2.5. Embryos micro-injection

For the micro-injection procedure, a mixture of sgRNA, zCas9 mRNA, and donor plasmids were co-injected into one-cell stage zebrafish embryos following fertilization. Each egg received an injection of 1 nl of solution, consisting of 600 ng/μl zCas9 mRNA, 100 ng/μl sgRNA, and 15 ng/μl donor plasmid.

2.6. Generation of transgenic zebrafish lines

Both the donor plasmid and the second intron of osr2 contain the sgRNA target site. Consequently, the simultaneous cleavage mediated by the sgRNA/Cas9 complex would facilitate precise and efficient integration of the donor DNA into the osr2 locus via the homologous recombination (HR) mechanism. Based on whether they expressed the green fluorescent protein cerulean in their eyes, four founders were identified for each transgenic line. These founders were crossed with WT zebrafish to obtain offspring. Subsequently, the extracted genomic DNA from these offspring served as the template to amplify the 5′ and 3′ junction fragments of targeted genes by the primers (F1: 5′-CGGTGACCGGACAGAGCAGACT-3′, R1:5′-ACTGGGTGCTCAGGTAGTGGTTGT-3’; 3.8 kb; F2:5′-GGAAGGCAAAATGCCGCAAAAAAGG-3′, R2:5′-AGCGGCCGCAGCAGAAAGGCTTGA-3’; 2.2 kb), and to sequence the 5′ junction using primer (F1: 5′-CGGTGACCGGACAGAGCAGACT-3’; R: 5′- ACACGCGCGCGCGCACTGAAC-3′) to identify potential indel mutations. Subsequently, the homozygous TgKI(osr2:GFF) lines was identified through sequencing and used in subsequent maintenance and experiments.

2.7. Zebrafish AKI model

The zebrafish AKI model was constructed through the intraperitoneal injection of gentamicin [28,29], following a previously described protocol [30]. In brief, adult Tg(osr2:EGFP) zebrafish lines of similar weight were subjected to intraperitoneal injection of gentamicin (2.7 μg/μL, 20 μL per fish). Subsequently, each injected adult zebrafish was individually placed in a separate container. Five to six adult zebrafish that produced similar amounts of visible solid proteinuria at 1 dpi (days post-injury) were used as experimental subjects for subsequent studies.

2.8. Nephron counting

Nephron counting was performed following a previously protocol [31]. Each nephron consists of only a segment of Tg(osr2:EGFP) PCTs. Therefore, quantification of nephron number relies on counting osr2:EGFP-labeled PCTs fragments. For counting, we carefully extracted kidneys from adult Tg(osr2:EGFP) fish and then performed whole-kidney imaging using a Nikon A1 confocal microscope. Subsequently, the total count of EGFP-positive segments was determined utilizing ImageJ. For unbiased analysis, all images were relabeled and counted by analysts without knowledge of each sample's treatment conditions. For evaluating variation in nephron number following Gentamicin-induced AKI, the baseline nephron count in uninjured kidneys was considered 100%. Subsequently, the extent of renal regeneration was characterized using the percentage change in nephrons after AKI compared to the uninjured state.

2.9. WISH (whole-mount in situ hybridization) of zebrafish embryos

The slc20a1a probe was synthesized (slc20a1a probe F: 5′-CATCGGAGGATCGGCAGAAACCGACC-3’; slc20a1a probe R: 5′-TCCGATGTTGGAAGCGACAACCACAG-3′) and WISH was performed following previously protocol [30]. Zebrafish embryos were fixed in 4% paraformaldehyde (PFA) overnight at 4 °C. Subsequently, the samples were washed five times with PBT (PBS with 0.1% Tween-20) and were permeabilized with proteinase K (10 μg/μL, Roche) in PBT for 5 min with rocking. The digoxigenin-labeled slc20a1a probe, synthesized from cDNA fragments containing zebrafish-specific sequences, was used for hybridization as previously reported [30]. Zebrafish embryos were subjected to hybridization with the digoxigenin-labeled slc20a1a probe. Anti-DIG AP antibody and NBT/BCIP substrate (Roche) were used to detect the probe. Following the color reaction, images were captured using a BX3-CBH microscope (Olympus, Japan).

2.10. Acquisition of EGFP signal

To analyze the expression pattern of EGFP, zebrafish embryos were treated with PTU (Sigma) (0.16 mM) from 16 h post-fertilization (hpf) to suppress pigmentation. At the indicated time, images were captured using both confocal microscopy and stereoscopic fluorescence microscopy to visualize the EGFP signal.

2.11. RT‒PCR and qRT‒PCR

RNA was extracted from kidney tissues utilizing TRIzol reagent (Invitrogen, 15596018). Subsequently, cDNA was synthesized from the extracted RNA using a Prime Script II 1st strand cDNA Synthesis Kit (Takara, 9767), followed by PCR using TB Green Premix EX TaqII (Takara, RR820A) for quantitative Real-time PCR (qRT‒PCR) (osr1-qRT: 5′-AGCATCCAGCAAAAGTAAACCC-3’; osr1-qRT: 5′-TTATGACAGATATCACAAGTGTACG-3’; 310 bp); β-actin [8]) or Taq Master Mix (Vazyme, p112-01) for reverse transcription polymerase chain reaction (RT‒PCR) (osr2-RT F: 5′- ATGGGCAGTAAGACTCTCCCGGCTCCGG-3’; osr2-RT R: 5′-CTAGACTGTGGCGCCGCAGGTCGCAGTT-3’; 710 bp; osr2-GFF-RT F: 5′-AAATCCTACAACCTGCTGATCCAC-3’; osr2-GFF-RT R: 5′-CCTGTTAACAATGCTTTTATATCCTGTA-3’; 560 bp). Gene expression data were normalized to the expression of β-actin mRNA.

2.12. Statistics

All experiments were performed in at least three independent replicates. Statistical analyses were conducted employing GraphPad Prism (version 10) and Excel (Microsoft Office Home and Student 2016 version). The data were analyzed by two-sided t-test and are presented as mean values ± S.

3. Results

3.1. Expression of osr2 in zebrafish PTECs

To investigate the expression pattern of osr2 in the zebrafish kidney, we conducted a reanalysis of previously published single-cell RNA sequencing (scRNA-seq) data (GSE100910) [26]. Analyzing scRNA-seq data revealed that osr2 is expressed in proximal tubule epithelial cells (PTECs) (Fig. 1A and B). Since the PT consists of the PCT and PST, we aimed to further validate the expression pattern of osr2 in the PT of the zebrafish kidney. To accomplish this, Seurat (version 4.2.0) was used to divide the PTECs cell population into subpopulations of PSTCs and PCTCs (Fig. 1C), based on marker genes that are specific and highly expressed in PCT (slc20a1a) and PST (slc13a1 and slc13a3) [27]. Notably, the analysis demonstrated that osr2 expression localized to the PCT of the zebrafish kidney (Fig. 1F). Moreover, the heatmap illustrated the expression of several PCT marker genes, including osr2, slc20a1a, slc6a19b, epdl2, slc7a7, slc34a1a, etc (Fig. 1D). Previous investigations have identified slc20a1a as a PCT marker [27], and our findings consistent with this observation, indicating that both slc20a1a and osr2 are expressed in the PCT (Fig. 1D–F). Collectively, these findings suggest that osr2 could serve as a valuable new marker for characterizing and studying the PCT.

Fig. 1.

Expression pattern of osr2 in zebrafish kidney. A-B: scRNA-seq analysis showed that osr2 was expressed in PTECs. tSNE plots show zebrafish kidney cell clusters and the expression of osr2. HSCs, hematopoietic stem cells; KPs, Kidney progenitors; KM, Kidney Multiciliated; MM, Macrophages Myeloid; KM, Kidney mucin; VECs, Vascular endothelium cells; DTECs, distal tubule epithelial cells; PTECs, proximal tubule epithelial cells. C: Cluster DTECs was divided into PSTCs and PCTCs subpopulations. D: Heatmap shows relative log-expression of some genes in PCT or PST in C. E-F: Expression analysis of slc20a1a and osr2 in each subpopulation. PCTCs, proximal convoluted tubule epithelial cells; PSTCs, proximal straight tubule epithelial cells.

3.2. PCT are marked by osr2:EGFP in zebrafish

To further determine whether osr2 can be used as a PCT marker,we designed a short guide RNA (sgRNA) targeting the second intron of zebrafish osr2. We then conducted co-injections of this sgRNA with zCas9 mRNA into one-cell-stage zebrafish embryos, resulting in a cleavage efficiency of ∼16%. Next, we constructed a donor plasmid osr2-P2A-GFF consisting of four parts: a left arm, a P2A-GFF coding sequence, a right arm, and crayy-EGFP (Fig. 2A). This approach facilitated knock-in of the whole donor plasmid fragment (Genomic location: GRCz11; Chromosome 16:54404070–54404071; Gene ID: ENSDARG00000038006) to generate TgKI(osr2:GFF) zebrafish (Fig. 2B;Fig. S1), which has been submitted the details to the zebrafish database ZFIN (the line ID: ZDB-ALT-231019-1). PCR and sequencing analysis of F1 progenies verified the inheritance of the genomic integration of their respective F0 founders, and TgKI(osr2:GFF) zebrafish with osr2⁻¹² (a 12-bp deletion in intron) was used for subsequent maintenance and experiments (Fig. 2C;Fig. S2). Subsequently, we examined osr2 transcription in TgKI(osr2:GFF) homozygous and WT using RT-PCR (Fig. 2D and E) and did not detect intact osr2 transcription in TgKI (osr2:GFF) homozygous. Therefore, the full reading frame of osr2 in TgKI (osr2:GFF) homozygous was disrupted. Given that GFF can activate transcription through its recognition sequence (upstream activating sequence, UAS), genes of placed downstream of UAS can be expressed in the GFF-expressing cells [24]. Tg(UAS:EGFP) transgenic reporter fish carries the EGFP gene downstream of UAS and can specifically express EGFP in GFF positive cells and tissues [24]. We crossed TgKI(osr2:GFF) with Tg(UAS:EGFP) to generate TgKI(osr2:GFF) x Tg(UAS:EGFP), abbreviated as Tg(osr2:EGFP), and observed EGFP expression in the pronephros of embryos (Fig. 2F and G). Based on their expression pattern and morphological characteristics, we postulated that the EGFP-expressing cells were localized within the PCTs. To further verify this, we employed WISH of slc20a1a, an established marker for PCTs [32], as a reference to validate the expression region of osr2:EGFP was the PCT. Our results showed that slc20a1a and osr2:EGFP were expressed at the same position, suggesting that osr2:EGFP marked the PCT (Fig. 2H–K), which was consistent with the results obtained from single-cell data analysis.

Fig. 2.

Intron targeting-mediated donor plasmid knockin at the zebrafish osr2 locus. A: Schematic representation of an intron-targeted mediated strategy for generating GFF knockin at zebrafish osr2 locus using the CRISPR/Cas9 system. The sequence of sgRNA target is shown in red, and the protospacer adjacent motif (PAM) sequence is depicted in green. The right and left arm sequences of the donor plasmid are denoted by the brown lines. The left arm is 840 bp, and the right arm is 840 bp. Following co-injection of the donor plasmid with the zCas9 mRNA and sgRNA, the whole donor plasmid fregment was integrated into the osr2 locus. The zebrafish osr2 has 4 exons, with E2, E3, and E4 representing the 2nd, 3rd, and 4th exons, respectively. B: PCR analysis of the 3′ and 5′ junctions of F1 progenies from the 3# founder. The F1, R1, F2 and R2 primers are shown in A. C: 5′ junction sequences of F1 progenies of four donor plasmid knockin F0 founders. The indel mutations are highlighted in red. D: Transcription schematic of osr2 in TgKI(osr2:GFF) and WT zebrafish. E: RT-PCR was used to verify the transcription of osr2 and GFF in TgKI (osr2:GFF) and WT zebrafish (n = 3 biological replications per group). F: Schematic diagram for obtaining TgKI(osr2:GFF) homozygotes, TgKI(osr2:GFF) heterozygotes, and Tg(osr2:EGFP) transgenic fish. G:In vivo confocal image (dorsal view) of Tg(osr2:EGFP)embryos at 24 hpf reveals EGFP expression in the pronephros as indicated by the red arrowheads. Scale bar, 500 μm. H–K: The expression patterns of slc20a1a and EGFP were examined in 24 hpf embryos. H: WISH analysis of the expression pattern of slc20a1a in 24 hpf embryos (side view). I:In vivo confocal images (side view) of osr2:EGFP embryos at 24 hpf. Posterior gut (white arrowhead). J: WISH analysis of the expression pattern of slc20a1a in 24 hpf embryos (dorsal view). K:in vivo confocal images (dorsal view) of Tg(osr2:EGFP) embryos at 24 hpf. hom, homozygotes; het, heterozygotes; pA, poly(A). Scale bar in H to K, 500 μm. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

3.3. Dynamic EGFP expression pattern during embryogenesis in Tg(osr2:EGFP) zebrafish

Expression of osr2 has previously been documented in various tissues of developing zebrafish embryos, including the pronephros, pectoral fin buds, and gut [[33], [34], [35]]. To assess whether Tg(osr2:EGFP) can faithfully reflect the expression of endogenous osr2, we performed a dynamic analysis of the osr2:EGFP expression pattern using stereoscopic fluorescence microscope. Through observation, we determined that osr2:EGFP was expressed in the PCT (Fig. 3A). As embryonic development progressed, the osr2:EGFP-labeled PCT showed a progressive aggregation and forward curling phenomenon (Fig. 3B–D). Additionally, osr2:EGFP expression was also detected in the pectoral fins, pharygial arch 1, and neuronal cells from 48 hpf to 120 hpf (Fig. 3A–D). Previous study has shown that osr2 is expressed in posterior gut epithelial cells from as early as 24 hpf, and in posterior gut and gut mesenchyme at 48 hpf [36]. To further validate osr2:EGFP is expressed in gut, we crossed Tg(osr2:EGFP) with Tg(cdh17:DsRed), which specifically marks the proximal and distal tubules of the pronephron. As a result, we observed that osr2:EGFP-expressing cells in the posterior gut were located below the renal tubules at 48 hpf and 72 hpf (Fig. 3E and F). As the embryo developed, posterior gut morphogenesis exhibited a gradual aggregation (Fig. 3G and H). Collectively, these findings demonstrate that Tg(osr2:EGFP) transgenic fish faithfully reflect the endogenous osr2 expression pattern.

Fig. 3.

Dynamic EGFP expression pattern throughout embryogenesis in the Tg(osr2:EGFP) transgenic line. A-D: Showing the PCT dorsal view of Tg(osr2:EGFP) embryos at 48 hpf, 72 hpf, 96 hpf, and 120 hpf, respectively. Within these stages, EGFP was expressed in the PCT region (indicated by the red arrowhead) as well as in the pectoral fin (indicated by the white arrowhead). E-F: Providing lateral view of Tg(osr2:EGFP;cdh17:DsRed) embryos at the developmental stages. Posterior gut (white arrowhead) and the gut lumen (*) can be observed. het, heterozygotes. Scale bar in A to H, 500 μm. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

3.4. Tg(osr2:EGFP) marks PCT morphogenesis

In order to further clarify Tg(osr2:EGFP) marks PCT morphogenesis, we employed confocal microscopy to examine the expression pattern of EGFP in the PCT. Our observations revealed that EGFP fluorescence signal in the developing kidney is initially detectable above the dorsal side at about 20 hpf. By 24 hpf, the EGFP fluorescence extends above the yolk sac, exhibiting bilateral tubular structures (Fig. 4A). To provide additional evidence for osr2 marking PCT morphogenesis, we crossed Tg(osr2:EGFP) with Tg(cdh17:DsRed). Subsequently, we tracked the expression patterns of osr2:EGFP and cdh17:DsRed in the embryos from 24 hpf to 120 hpf (Fig. 4A–E). Our results demonstrated co-expression of EGFP and DsRed in the PCT throughout embryonic development (Fig. 4) and persisting into adulthood. The temporal progression of slc20a1a-expressing PCT morphogenesis, transitioning from a linear tube to a coiled structure, had been previously characterized [27]. Our observed temporal progression of PCT morphogenesis, marked by osr2:EGFP, is consistent with these findings. Therefore, based on the results of single-cell analysis, in situ hybridization, and transgenic fish, we believe that osr2 is a new marker gene for PCT.

Fig. 4.

The PCT morphogenesis is labeled by EGFP in Tg(osr2:EGFP) embryos. A-E: Dorsal view of the PCT cells that are co-labeled by EGFP and DsRed at the indicated stages are shown (n = 6). osr2:EGFP expression in the pectoral fins (red arrowhead) can be observed. Scale bar in A to E, 200 μm. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

3.5. osr2 mutants do not affect zebrafish PCT development

Previous studies on the role of osr2 in zebrafish kidney development have yielded conflicting results. Therefore, it is crucial to gather additional evidence to determine the impact of osr2. In our knock-in strategy, the whole donor plasmid was integrated into zebrafish genomic (Fig. 2A and B;Fig. S1), resulting in the full reading frame of osr2 in TgKI (osr2:GFF) homozygous was disrupted (Fig. 2D and E). Consequently, TgKI(osr2:GFF) homozygotes are mutants, while the above developmental studies employed TgKI(osr2:GFF) heterozygotes that do not exhibit any discernible effects on embryo development. To investigate whether osr2 mutants affect embryogenesis, we monitored the growth of Tg(osr2:EGFP;cdh17:DsRed) zebrafish embryos using a stereomicroscope. As a result, the morphology and development of PCT and renal tubules in osr2-inactivated Tg(osr2:EGFP;cdh17:DsRed) zebrafish embryos were indistinguishable from the control group (Fig. 5A–J). Additionally, obvious deficiencies were not found in other organizations (pharygial arch 1, pectoral fins, and neuronal cells) in TgKI(osr2:GFF) homozygotes zebrafish. We believe that osr2 inactivation has no impact on the development of nephrons, possibly related to complementation of osr2 paralogous genes. In addition, we also detected the transcript level of osr2 paralogous, osr1, in embryonic and adult TgKI (osr2:GFF) homozygous and WT zebrafish kidneys by qRT-PCR, however, no obvious upregulation of osr1 mRNA was found (Fig. 5K). Collectively, we believe that osr2 inactivation does not seem to exert any discernible effects on zebrafish kidney development.

Fig. 5.

Embryo morphology of TgKI(osr2:GFF) homozygotes and TgKI(osr2:GFF) heterozygotes. A-J: Use of Tg(osr2:EGFP;cdh17:DsRed) background to show TgKI(osr2:GFF) heterozygotes (A-E) and TgKI(osr2:GFF) homozygotes (F-J) embryo developmental morphology, development of PCT and renal tubules at the indicated stages are shown (n = 6). K: qRT‒PCR analyses of osr1 in embryonic and adult TgKI(osr2:GFF) homozygous and WT zebrafish kidneys (n = 3 biological replications per group). hom, homozygotes; het, heterozygotes. Scale bar in A to J, 500 μm. The data in K were analyzed by two-sided t-test and are presented as mean values ± SD.

3.6. osr2:EGFP adult zebrafish provides a valuable tool for detection nephron regeneration

Fish kidneys have the remarkable ability to regenerate nephrons de novo after injury [30,37], designated renal regeneration. In response to injury, renal progenitor cells (RPCs) undergo a process of aggregation, followed by rapid proliferation and subsequent differentiation into new nephrons, providing the basis for new nephron regeneration count [8]. Despite the significance of nephron regeneration, there remains a lack of convenient and effective methods for nephron regeneration counting due to the inherent complexity of nephron structures. Our results indicated that osr2:EGFP was expressed in the PCT throughout embryonic development (Fig. 4) and persists into adulthood (Fig. 6B and C). Given that each nephron contains a single PCT segment, evaluating the regeneration of PCT segments enables an approximation of nephron regeneration rate. The labeled fragments observed in Tg(osr2:EGFP) transgenic fish are short and exhibit rarely overlap, thereby providing a useful tool for detecting the rate of nephron regeneration. Therefore, the use of Tg(osr2:EGFP) zebrafish greatly facilitates nephron regeneration counting. Subsequently, we aimed to determine the potential effects of osr2 inactivation on nephron regeneration in Tg(osr2:EGFP) zebrafish. To achieve this, we induced AKI by intraperitoneally injecting adult zebrafish with a well-established nephrotoxin, gentamicin (Gent; 2.7 μg/μL, 20 μL per fish) [28], according to the schematic diagram outlining the construction of the AKI model (Fig. 6A). Our findings showed that, in comparison to uninjured kidneys, both TgKI(osr2:GFF) homozygotes and TgKI(osr2:GFF) heterozygotes exhibited similar rates of nephron regeneration at 4, 6, and 8 days post-injury (dpi), approximately 18%, 50%, and 80%, respectively (Fig. 6D–L). Therefore, we conclude that the inactivation of osr2 may not affect nephron regeneration in Tg(osr2:EGFP) zebrafish compared to the control group.

Fig. 6.

EGFP ⁺ cells regeneration in Tg(osr2:EGFP) adult zebrafish after kidney injury. A: Schematic diagram of the AKI model constructed by intraperitoneal injection of gentamicin in adult zebrafish. B: Confocal images of adult zebrafish PCT cells that are co-labeled by osr2:EGFP and cdh17:DsRed are shown (n = 6). Scale bar in A, 100 μm. C: Higher-magnification image of the boxed area showed in B. Scale bar, 200 μm. D-G: Use of osr2:EGFP background to show of PCTs in TgKI(osr2:GFF) homozygotes at the indicated time point after injured (n = 6). Scale bar, 100 μm. H–K: Use of osr2:EGFP background to show of PCTs in TgKI(osr2:GFF) heterozygotes at the indicated time point after injured (n = 6). Scale bar, 100 μm. L: After injured, the renal regeneration rate of TgKI(osr2:GFF) homozygotes and TgKI(osr2:GFF) heterozygotes were counted using ImageJ (n = 5 biological replications per group). hom, homozygotes; het, heterozygotes. Scale bar in A to J, 500 μm.

4. Discussion

Transgenic technology offers numerous advantages, such as facilitating the integration of fragments containing fluorescent proteins and target genes, thereby facilitating early screening of experimental animals. In this study, we employed CRISPR/Cas9 knock-in technology to generate Tg(osr2:EGFP) zebrafish. Our findings revealed EGFP expression in the kidney, pectoral fin, and gut, exhibiting a pattern consistent with the endogenous osr2 expression profile. Consequently, Tg(osr2:EGFP) can faithfully reflect the expression of osr2 in vivo. By tracking the expression of EGFP, we can investigate the development of PCT. Thus, the establishment of Tg(osr2:EGFP) transgenic zebrafish lines laid a crucial foundation for our subsequent investigations on the development of PCT in zebrafish.

The PCT of kidney plays a crucial role in the secretion, reabsorption, and maintenance of electrolyte and water balance within the renal tubule. However, transgenic fish lines that mark the PCT development are still lacking. Two fluorescently labeled transgenes, Tg(cdh17:EGFP)^pt305 and Tg(enpep:GFP), driven by kidney-specific promoters (cdh17 and enpep) can mark the PT, but these transgenes also exhibit fluorescence in other kidney tubule cells [38,39]. Transgenic zebrafish line, Tg(gtshβ:GFP), driven by the grouper gtshβ promoter, expresses GFP in the pituitary and kidney, and marks the developing details of the PCT and PST segments [12]. However, an endogenous zebrafish line that clearly marks the PCT is still lacking. In this study, through reanalysis of previously published single-cell RNA sequencing (scRNA-seq) data (GSE100910), we found that osr2 was a novel mark for labeling the PCT (Fig. 1). osr2 has been reported to have a role in kidney development. Additionally, osr2 is expressed in the kidney earlier than slc20a1a [22,40], which can better track the PCT morphogenesis. Therefore, the establishment of Tg(osr2:EGFP) transgenic zebrafish line offers a valuable tool for tracking PCT morphogenesis.

Nephrons are highly complex structures, and their close proximity makes nephron counting challenging. Previous methods, such as in situ hybridization, have been utilized for nephron counting [8], but they are complicated and time-consuming. Another approach involves the use of 40 kDa dextran-FITC, which can be taken up by duct cells to count nephrons. However, the labeled fragments are too long, making nephron counting difficult. In contrast, the Tg(osr2:EGFP) transgenic fish, which labels the PCT fragment, exhibits shorter and less overlapping fragments, making it highly suitable for nephron counting.

To generate the TgKI(osr2:GFF) zebrafish line, the donor plasmid was inserted into zebrafish genome (Genomic location: GRCz11; Chromosome 16:54404070–54404071; Gene ID: ENSDARG00000038006) using CRISPR/Cas9 technology (Fig. 2A–E). This lines were then crossed with Tg(UAS:EGFP) to obtain the Tg(osr2:EGFP) line (Fig. 2F and G). Notably, the disruption of the full reading frame of osr2 resulted in the inactivation of osr2. Nevertheless, our study revealed no evident abnormalities in osr2-inactivated TgKI(osr2:GFF) zebrafish embryos development (Fig. 5). To investigate the impact of osr2 inactivation on nephron regeneration, we induced AKI in adult zebrafish through intraperitoneal injection of gentamicin. Nephron regeneration was subsequently assessed. As a result, the inactivation of osr2 did not affect nephron regeneration in adult zebrafish (Fig. 6). Based on these findings, we conclude that osr2 inactivation may not significantly impact zebrafish PCT development and regeneration.

Herein, both TgKI(osr2:GFF) homozygotes and TgKI(osr2:GFF) heterozygotes could grow normally to adulthood and reproduce normally. No obvious deficiencies were observed in zebrafish with inactivated osr2; however, the data from Morpholino experiments present conflicting outcomes, with some studies indicating the presence of defects [33], while others report no such abnormalities [36]. Consequently, we need to exclude the side effects of Morpholino. Additionally, genetic compensation response occurs when a gene mutation results in complete loss of function, prompting the body to increase the expression of other genes to replace its function for normal development and survival [41]. The genetic compensation response triggered by gene-knockout mutations in diverse organisms, such as zebrafish [42,43], mice [44] or Arabidopsis [45], were found to be a mechanism contributing to genetic robustness. In order to determine whether the loss of function of osr2 did not affect the growth of zebrafish due to genetic compensation, we detected the transcript level of the osr2 paralogous gene osr1, and found that the transcript level of osr1 did not change (Fig. 5K). Neither the side effects of Morpholino nor the influence of genetic compensation response can be excluded in our results. Therefore, whether osr2 affects kidney development requires detailed follow-up research.

In this study, we successfully generated Tg(osr2:EGFP) zebrafish using CRISPR/Cas9 knock-in technology. The Tg(osr2:EGFP) zebrafish line serves as a valuable model to observe the dynamic progression of PCT development throughout the entire lifespan through the tracing of EGFP signaling. Moreover, this transgenic fish line enables quantification of nephron regeneration. Overall, the Tg(osr2:EGFP) zebrafish line is an excellent tool for screening mutants associated with PCT development, as well as for screening potential drugs targeting PCT development and related diseases.

Data availability

No data was used for the research described in the article.

CRediT authorship contribution statement

Wenmin Yang: Writing – review & editing. Xiaoliang Liu: Writing – review & editing, Software. Zhongwei He: Software. Yunfeng Zhang: Writing – original draft. Xiaoqin Tan: Writing – original draft. Chi Liu: Writing – review & editing.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

The following is the Supplementary data to this article: