Generation of Mice Harboring Bicc1 Conditional Null Alleles

ABSTRACT

Bicaudal C1 (Bicc1) encodes an RNA‐binding protein critical for many organ development and epithelial tissue homeostasis. Bicc1 null mutations have been shown to lead to the development of polycystic kidney disease (PKD) and death at an early prenatal stage. To elucidate the tissue‐specific functions of Bicc1, we engineered two independent conditional knockout (cKO) mouse lines targeting distinct exonic regions of the gene. The first line was generated using a traditional embryonic stem (ES) cell‐based approach, wherein loxP sites were inserted flanking exon 4 (E4), enabling Cre‐mediated excision of a functionally essential coding region. The second line was created using CRISPR/Cas9 genome editing, introducing loxP sites around both exon 4 and exon 5 (E4‐5) in a double‐step zygote injection strategy. Both alleles were validated by PCR genotyping, sequencing, and functional recombination was confirmed via a tissue‐specific Cre driver. These independent cKO models provide a robust platform for dissecting the role of Bicc1 in specific tissues and developmental stages, and offer new avenues for studying the mechanistic basis of PKD and other Bicc1‐related pathologies.

1. Introduction

The evolutionarily conserved KH‐domain protein Bicaudal C homolog 1 (BICC1) is an RNA‐binding protein involved in cell fate determination, vertebrate organogenesis and tissue homeostasis (Dowdle et al. 2022; Park et al. 2016; Pedrozo et al. 2015; Saffman et al. 1998; Tran et al. 2007; Wessely and De Robertis 2000). Bicc1 contains three Kunitz Homology (KH) RNA‐binding domains (KH1–KH3) and two KH‐like domains (KHL1, KHL2). KH3 is flanked by KHL1 and KHL2, linked by a serine–glycine–rich sequence, and lies near the C‐terminus adjacent to the SAM domain (Mahone et al. 1995; Wessely and De Robertis 2000; Wessely et al. 2001). Through selective binding to target transcripts and recruitment of other SAM domain proteins, BICC1 regulates gene‐expression programs during development and in adult tissues (Rothé et al. 2015; Wessely and De Robertis 2000). Dysregulation of BICC1 has been linked to defects in left–right patterning, heart and renal morphogenesis, and epithelial integrity (Bouvrette et al. 2010; Maisonneuve et al. 2009; Rothé et al. 2020; Tran et al. 2010; Wessely and De Robertis 2000). Two Bicc1 knockout (KO) mouse lines and two spontaneous alleles (jcpk and bpk) have been reported (Cogswell et al. 2003; Maisonneuve et al. 2009; Tran et al. 2010). The two Bicc1‐KO mouse lines exhibit congenital heart defects, heterotaxy, and polycystic kidney disease (PKD) and die during early postnatal stages. In contrast, the jcpk and bpk mouse models develop PKD but no situs abnormalities have been reported (Rothé et al. 2020). Despite substantial progress, the in vivo roles of Bicc1, in particular, in the adult remain incompletely defined. While the conventional loss‐of‐function alleles have yielded valuable insights, their interpretation is confounded by early lethality and the inability to delineate tissue‐specific requirements. Thus, a conditional allele of Bicc1 that enables spatially and temporally controlled gene inactivation is essential to dissect the role of Bicc1 across organs and developmental stages.

To meet this need, we generated two independent conditional Bicc1 mouse lines in which loxP sites were placed in cis to flank a critical coding region encoding the RNA‐binding KH domains. Cre‐mediated excision removes exon 4 (E4) in one line (Bicc1 ^E4fl/E4fl ) and exons 4–5 (E4–5) in the other (Bicc1 ^E45fl/E45fl ), both of which encode a portion of the KH1 domain (with exons 4–5 additionally encompassing KH2), resulting in frameshift alleles expected to trigger nonsense‐mediated decay. We deliberately employed complementary engineering strategies to ensure robustness and facilitate cross‐validation: (i) a CRISPR/Cas9‐assisted genome‐editing approach and (ii) a classical embryonic stem (ES) cell–targeting approach. These parallel strategies minimize the risk of line‐specific artifacts, allow independent haplotype backgrounds, and provide orthogonal routes to reproducibility. The unrecombined floxed alleles were designed to be phenotypically silent and each allele was molecularly validated by junction PCR and Sanger sequencing.

As a functional benchmark, we combined each floxed allele with a kidney‐epithelial specific Cre driver (Cdh16‐cre) to generate kidney‐specific Bicc1 null animals. In both lineages, tissue‐specific deletion resulted in early postnatal lethality and PKD phenotypes, consistent with an essential role for Bicc1 in neonatal viability and providing strong evidence that Cre‐mediated recombination of our conditional alleles yields a true loss‐of‐function state.

Together, these conditional alleles of Bicc1 establish a versatile platform for spatiotemporal knockout studies in vivo. By enabling cell‐type–specific and stage‐specific gene ablation, they will allow the field to define upstream regulators and downstream effectors of Bicc1 in discrete physiological contexts, to map tissue susceptibilities, and to separate developmental from homeostatic functions. The availability of two independently engineered lines will provide robustness, an internal control for phenotype reproducibility, and will support rigorous genetic epistasis analyses.

2. Results and Discussion

We generated conditional Bicc1 knockout alleles using two strategies, classical embryonic stem (ES) cell targeting and a CRISPR/Cas9‐based approach. For the ES cell approach, a targeting construct was built on a C57BL/6 background. Two loxP sites were positioned to flank E4, and the sites were separated by ~1.6 kb to enable efficient Cre‐mediated excision (Figure 1A). E4 encodes part of the RNA‐binding domain KH1 region, and deletion thereof introduces a frameshift that is predicted to trigger nonsense‐mediated decay of the primary transcript. The targeting vector also carried an FRT‐flanked neomycin cassette for positive selection upstream of the 3′ loxP site and a thymidine kinase cassette for negative selection. Following electroporation and selection with neomycin/ganciclovir, three correctly targeted ES clones were recovered; two of the clones were used to produce male chimeras, of which ultimately one line with robust germline transmission was maintained. Germline transmission was established by breeding chimeric males with B6 albino females (B6N‐Tyr^c‐Brd) and identifying transmission to offspring by black color coat. The selection cassette was then deleted by crossing to hACTB‐FLPe transgenic mice to delete the Neo cassette producing Bicc1 ^E4fl/+ animals; subsequently, mice were outcrossed to C57B6 wild‐type mice to remove the FLPe transgene (Figure 1A). Homozygous floxed mice were generated by intercrossing Bicc1 ^E4fl/+ males and females yielding Bicc1 ^+/+ , Bicc1 ^E4fl/+ , and Bicc1 ^E4fl/E4fl progeny. Genotyping PCR identified Bicc1 ^E4flE4/fl offspring (Figure 1B). Sanger sequencing across both loxP insertions verified an intact floxed configuration without unintended rearrangements in the Bicc1 locus (Figure 1C).

FIGURE 1.

Generation of a conditional Bicc1 allele by ES cell targeting. (A) Schematic of the targeting strategy. E4 of Bicc1 was flanked by two loxP sites; an FRT‐flanked neomycin cassette (Neo) was positioned upstream of the 3′ loxP site for positive selection, and a thymidine kinase (TK) cassette was included for negative selection. Flpe‐mediated excision of the frt‐Neo‐frt‐loxP cassette from Bicc1 ^f/lNEO allele resulted in the Bicc1 ^E4fl/+ allele. Cre recombination deletes E4 containing part of KH1 creating a null allele. (B) PCR genotyping strategy. Left: PCR across the 5′ loxP site (primers P1–P2) amplified a 665 bp fragment from the targeted allele and a 631 bp fragment from wild type. Right: PCR across the 3′ loxP site (primers P3–P4) amplified a 414 bp fragment only from the floxed allele and no product from the wild‐type allele. Representative genotyping results are shown. (C) Sequence validation of loxP insertions. Sanger sequencing of PCR products spanning each insertion in Bicc1 ^E4fl/E4fl mice confirms the canonical 34‐bp loxP sequence (highlighted in blue) in its correct orientation at both the 5′ and 3′ sites and no evidence for any unintended genomic rearrangements.

For the CRISPR/Cas9‐based approach, we installed two loxP sites flanking the Bicc1 E4 and E5 interval using a same‐day/two‐step strategy. The two loxP sites were 3.4 kb apart, which is still within the optimal distance of cre recombination efficiency. Guide RNAs targeted intronic sequences upstream of E4 and downstream of E5 (Figure 2A). Cas9 ribonucleoprotein complexes, guide RNAs, and single‐stranded donor oligonucleotides bearing a 34‐bp loxP sequence with short homology arms were electroporated into zygotes. The 5′ loxP site was introduced in the morning; after a brief recovery, the 3′ loxP site was introduced into the same embryo cohort in the afternoon. Three out of nine founders were obtained. PCR genotyping and Sanger sequencing confirmed the correctly targeted allele with two intact loxP sites in cis on the same chromosome. Because founders are mosaic, we bred the founders to C57BL/6 wild‐type mice to generate F1 offspring with germline transmission carrying the Bicc1 ^E45fl/+ allele. We then interbred Bicc1 ^E45fl/+ F1 animals to obtain Bicc1 ^E45fl/E45fl homozygous mice. These mice were screened by junction PCR using primer pairs flanking each insertion site (Figure 2B). Amplicons showing the expected size shift were Sanger sequenced to verify the intact loxP sequence at both the 5′ and 3′ sites (Figure 2C). These Bicc1 ^fl/fl animals, carrying two intact loxP sites flanking E4, or E4–E5, served as the validated conditional line for subsequent Cre‐dependent studies.

FIGURE 2.

CRISPR/Cas9‐mediated generation of a floxed Bicc1 allele spanning E4 to E5. (A) Schematic of the editing strategy. Two loxP sites were inserted into intronic regions flanking E4 and E5, separated by ~3.4 kb. Ultramer donor oligonucleotides were used to introduce the canonical 34 bp loxP sequence (red) along with engineered PvuII and NcoI restriction sites for screening. Guide RNA target sequences are shown in bold and PAM sequences in green. (B) F1 genotyping by PCR. PCR using primer pairs P5‐P6 and P7‐P8 confirmed insertion of the 5′ and 3′ loxP sites. The targeted alleles generated 328 bp amplicons at the 5′ end and 358 bp at the 3′ end, compared to amplicons of 290 bp at the 5′ and 321 bp at the 3′ in wild type. Representative PCR results are shown. (C) Sanger sequencing of PCR amplicons verified the presence and correct orientation of the canonical loxP sequence (highlighted in blue) at both the 5′ and 3′ sites.

Both Bicc1 ^E4fl/E4fl and Bicc1 ^E45fl/E45fl mice exhibited body weights within the expected range for C57BL/6J wild‐type controls (JAX 2005), and both sexes were fertile (Figure 3). This indicates that the introduction of loxP sites flanking E4 or E4 and E5 in the absence of a Cre recombinase did not affect viability or overall embryonic development.

FIGURE 3.

Body weight analysis of floxed Bicc1 mouse lines. (A, B) Growth curves and body weight tables for Bicc1 ^E4fl/E4fl (A) and Bicc1 ^E45f/lE45fl mice (B). Male (blue) and female (pink) mice were weighed weekly from 4 to 12 weeks of age. Data are presented as mean ± SD. Tabulated values below the graph show weekly means with standard deviations with an N = 6–11 in (A) and an N = 8–11 in (B). Body weights were within the expected range for strain‐matched C57BL/6 wild‐type mice.

To confirm Cre‐mediated recombination in vivo, we crossed Bicc1 ^E4fl/E4fl or Bicc1 ^E45fl/E45fl mice with Bicc1 ^+/− mice (Tran et al. 2010) harboring a Cdh16‐cre transgene (Bicc1 ^+/− ; Cdh16‐cre) (Figure 4A), which drives recombination in epithelial cells of the developing kidney and genitourinary tract, as well as in the renal tubules of adult mice (Shao et al. 2002). Both Bicc1 ^E4fl/− ; Cdh16‐cre and Bicc1 ^E45fl/− ; Cdh16‐cre conditional knockout (cKO) mice were born in a normal Mendelian ratio. However, they developed bilateral nephromegaly with a severe polycystic phenotype (Figure 4B,C) and died between postnatal day (P) P10 and P20, likely due to renal failure. The PKD phenotypes mirror those reported for Bicc1‐null mutants (Maisonneuve et al. 2009; Tran et al. 2010). Importantly, neither cKO line exhibited heterotaxia, a defect that arises during early embryogenesis (Maisonneuve et al. 2009; Tran et al. 2010). The absence of left–right patterning abnormalities confirms the expected tissue specificity of the Cdh16‐cre transgene.

FIGURE 4.

Kidney epithelial deletion of Bicc1 results in a PKD phenotype. (A) Experimental scheme for generating the nephron epithelial conditional Bicc1 mice. (B) Gross kidney morphology. Both Bicc1 ^E4fl/− ; Cdh16‐cre (n = 12) and Bicc1 ^E45fl/− ; Cdh16‐cre (n = 8) kidneys display severe cystic dilation and loss of cortical‐medullary distinction when compared to the kidneys of control mice (n = 22 for Bicc1E4 conditional knockout controls, n = 12 for Bicc1E45 conditional knockout controls). Scale bars = 1 mm. (C) Hematoxylin and eosin (H&E)‐stained kidney sections reveal normal parenchyma in controls, but extensive cyst formation, thinning of renal tissue, and disrupted architecture in both conditional knockout lines. Scale bars = 1 mm. (D) Determination of cyst origin by immunofluorescence staining. Staining with segment‐specific markers demonstrates loss of tubular organization in cystic kidneys LTA (yellow), and DAPI (blue) mark distal tubules (top row). THP (orange), and DAPI (blue) identify thick ascending limb (TAL) (second row). LEA (yellow), and DAPI (blue) mark distal tubules (third row). AQP2 (orange), LTA, and DAPI (blue) label collecting ducts (fourth row). NCC (orange) and DAPI (blue) label the distal convoluted tubule. Control kidneys show well‐organized, segment‐specific staining, whereas Bicc1 ^E4fl/− ; Cdh16‐cre and Bicc1 ^E45fl/− ; Cdh16‐cre kidneys exhibit severely distorted tubule morphology and cystic dilation in the TAL, distal tubules and the collecting duct. Please note that the upper left control image and the upper right Bicc1 ^E45fl/− ; Cdh16‐cre image in Figures 4 and 5 are consecutive tissue sections. Control mice were littermates without a homozygous Bicc1 deletion (Cdh16‐cre; Bicc1 ^E4fl/+ , Bicc1 ^E4fl/+ ; Cdh16‐cre, Bicc1 ^E45fl/ , or Bicc1 ^E45fl/+ ; Cdh16‐cre). Scale bars = 100 μm.

To define the segmental origin of the cysts, we performed immunohistochemistry on kidneys from control and cKO littermates using markers for specific nephron segments, Lotus tetragonolobus agglutinin (LTA, proximal tubule), Tamm–Horsfall protein/uromodulin (THP, thick ascending limb or TAL), the thiazide‐sensitive Na⁺‐Cl⁻ cotransporter (NCC, distal convoluted tubule), Lycopersicon esculentum (LEA, distal tubule), and aquaporin‐2 (AQP2, collecting duct). LTA‐positive cysts, indicative of a proximal tubule origin, were rare in the cKO kidneys (Figure 4D, top row). In contrast, cyst‐lining epithelia were frequently positive for THP, LEA, and AQP2 (Figure 4D). Additionally, we observed the rare cysts were positive for NCC. This indicates that most cysts arise from the TAL, distal tubule and the collecting duct, with only a limited contribution from the proximal tubule. These findings correlate with the reported Cdh16‐cre expression, which shows that Cre recombinase activity is present in the TAL, distal tubules and cortical collecting ducts, but mostly absent in proximal tubules (Shao et al. 2002). It also demonstrates the benefit of the conditional Bicc1 allele, as global Bicc1 mutants exhibit cysts in the glomerulus and proximal tubules (Tran et al. 2010). In the future, it will be interesting to further dissect the segment‐specific or developmental timing effects by eliminating Bicc1 using Cre driver lines that, for example, specifically target proximal tubular cells or can be activated postnatally.

Using immunofluorescence for Ki67, a marker of cycling cells, we observed an increase in proliferating cells in both Bicc1 ^E4fl/− ; Cdh16‐cre and Bicc1 ^E45fl/− ; Cdh16‐cre kidneys compared with controls (Figure 5). In control kidneys, Ki67+ nuclei were sparse and scattered throughout the kidney (Figure 5A1–A3,D10–D12). In the cKO mutant kidneys, Ki67+ nuclei were much more prominent (Figure 5B,C,E,F). Numerous Ki67‐positive cells localized to LEA‐positive cyst walls, indicating robust cell‐cycle entry within distal tubular–derived cyst epithelia (Figure 5E13–E15,F16–F18). Based on comparison with THP staining in adjacent sections, Ki67‐positive cells were also present in regions corresponding to the TAL. Yet, double staining of Ki67 and LTA revealed very few double‐labeled cells (Figure 5B4–B6). Together, these data demonstrate that proliferation associated with Bicc1 loss occurs predominantly in non‐proximal epithelia and likely contributes to cyst expansion. Of note, immunostaining with nephron segment–specific markers was used to determine cyst origin. However, this approach was not intended to quantify marker protein expression in the cKO kidneys. Because the severely cystic kidneys exhibit marked structural distortion, the arrangement of nephron segments in thin sections often varies. As such, any apparent differences in staining patterns are more likely attributable to differences in tissue architecture rather than underlying biology. Future quantitative approaches, such as confocal 3D imaging or isolation of specific nephron segments for molecular analysis, are required to precisely assess potential segment‐specific effects of the two alleles.

FIGURE 5.

Increased proliferation in cystic kidneys of conditional Bicc1 knockout mice.Immunofluorescence staining of kidneys from control mice (A, D) and Bicc1 ^E4fl/− ; Cdh16‐cre (B, E) or Bicc1 ^E45fl/− ; Cdh16‐cre (C, F) conditional knockouts. Sections were stained for Ki67 (magenta), LTA or LEA (yellow), and DAPI (blue). (A–C) Ki67/LTA/DAPI staining shows sparse Ki67+ nuclei in control kidneys (A), whereas numerous Ki67+ cells are present in cyst‐lining epithelia of both cKO models (B, C). (D–F) Ki67/LEA/DAPI staining reveals abundant proliferating cells within LEA‐positive cyst walls in cKO kidneys (E, F), while control kidneys show only scattered Ki67+ cells (D). Insets correspond to higher‐magnification views of the boxed regions: Panels 1–3 from A; 4–6 from B; 7–9 from C; 10–12 from D; 13–15 from E; and 16–18 from F. Please note that the upper left control image and the upper right Bicc1 ^E45fl/− ; Cdh16‐cre image in Figures 4 and 5 are consecutive tissue sections. Control mice were littermates without a homozygous Bicc1 deletion (Cdh16‐cre; Bicc1 ^E4fl/+ , Bicc1 ^E4fl/+ ; Cdh16‐cre, Bicc1 ^E45fl/+ or Bicc1 ^E45fl/+ ; Cdh16‐cre). Scale bars = 100 μm.

In conclusion, the two independently engineered conditional‐null alleles of Bicc1 reveal a consistent, rapidly progressive PKD phenotype driven by hyperproliferation in the distal nephron and collecting duct epithelia, which are the cells targeted by the Cdh16‐cre. This provides a robust platform to dissect Bicc1 function across developmental and early postnatal stages. Because the Cdh16‐cre is expressed as early as E15.5, recombination occurs in the embryonic kidney explaining the early postnatal lethality and preventing the assessment of adult kidney function. As a future direction, postnatal, inducible renal epithelial Cre drivers, such as Cdh16‐creER ^T2 (Patel et al. 2008) after weaning or in the adult, and potentially segment‐restricted lines (e.g., Aqp2‐CreER^T2 or Umod‐CreER^T2) could be used to inactivate Bicc1 in the mature kidney, thereby disentangling kidney development from cyst initiation/progression.

3. Material and Method

3.1. Gene Targeting and Generation of Mutant Mice

The Bicc1^E4fl‐neo targeting vector and ES cell clones were generated by the Genetically Engineered Models Core at Louisiana State University. CRISPR targeting reagents were designed and produced by the Transgenic Animal and Genome Editing (TAGE) Core at Cincinnati Children's Hospital Medical Center. Briefly, the Bicc1 ^E45fl/+ conditional knockout allele was generated via a same‐day sequential loxP insertion approach (Figure 2A). Briefly, the 5′ and 3′ single‐guide RNAs with the target sequence of 5′‐CCC AGA CCT TGG TGC AGT GA‐3′ and 5′‐CCG CAG CTT TAT CCG GTT GG‐3′, respectively, were selected according to their on‐ and off‐target scores from the web tool CRISPOR (http://crispor.tefor.net) and synthesized from IDT. Similarly, the donor oligo was designed using an asymmetrical homologous arm design containing the loxP sequence and synthesized as an Ultramer from IDT. To form the ribonucleoprotein complex (RNP), individual sgRNA (60 ng/μL) was mixed with Cas9 protein (IDT; 80 ng/μL) in Opti‐MEM (ThermoFisher) and incubated at 37°C for 15 min. The donor oligo was added to the two RNPs at the final concentration of 475 ng/μL. The zygotes from super‐ovulated female mice on the C57BL/6J background were first electroporated with 7 μL RNP/donor mix targeting the 5′ site using a Genome Editor electroporator (BEX; 30 V, 1 ms width, and 5 pulses with 1 s intervals). Two minutes after electroporation, zygotes were transferred into 500 μL cold M2 medium (Sigma), brought to room temperature, and then cultured in KSOM (CytoSpring) containing 2 μM of the DNA activated protein kinase inhibitor M3814 (AOBIOUS) in a 5% CO2 incubator at 37°C. Four hours later, the zygotes were electroporated again now targeting the 3′ site. Zygotes were subsequently transferred into the oviductal ampulla of pseudopregnant CD‐1 females. To increase the efficiency of producing homozygous Bicc1 null alleles and decrease the incidence of mosaic deletion, we generated Bicc1 ^E4fl/− ; Cdh16‐cre and Bicc1 ^E‐5fl/− ; Cdh16‐cre mice. These mice were derived from breeding of Bicc1 ^+/− ; Cdh16‐cre and Bicc1 ^E4fl/E4fl or Bicc1 ^E45fl/E45fl . The Bicc1 ^+/− mice were generated previously (Tran et al. 2010). The offspring carrying 5′ and 3′ loxP in cis were identified by PCR and Sanger sequencing. The hACTB‐FLPe transgenic mice and Cdh16‐cre were obtained from Jackson Laboratories. The B6N‐Tyrc‐^Brd albino female mice were from Charles River. Animals were housed in a controlled environment with a 12 h light/12 h dark cycle, with free access to water and a standard chow diet. All animal procedures were carried out in accordance with the Institutional Animal Care and Use Committee‐approved protocol of the Cleveland Clinic Foundation and Cincinnati Children's Hospital and Medical Center. The Bicc1 ^flfl animals will be made available to the research community upon acceptance of the manuscript.

3.2. Genotyping

Genomic DNA was isolated from mouse tissues and analyzed by PCR using standard protocols; primer sequences were as follows: P1, 5′‐ACT ATT GTT GCT TTC TGC ATT GG‐3′; P2, 5′‐CCT TTC AAA CTT TGC CGA TTT CT‐3′; P3, 5′‐CAG TCA GGT ACA TAA TAT AAC TTC GTA TAA TG‐3′; P4, 5′‐CCT GAA CAG GAA GAC TCA AGA TAT AA‐3′; P5, 5′‐GTC TTG TAT CCC ACG CTC TGT C‐3′; P6, 5′‐CAC AGA ACA ACA AAG TCC ACT TGG‐3′; P7, 5′‐GTA CTC TGT GGG ACC AAG GAT G‐3′; P8, 5′‐TCA TGG GAG CAG CTA AGA AGA C‐3′.

3.3. Histology Analysis and Immunohistochemistry

The kidneys were fixed in 4% paraformaldehyde at 4°C overnight on a rocking platform, then rinsed in PBS, dehydrated through a graded ethanol series, cleared, and embedded in paraffin. Paraffin blocks were sectioned on a microtome at 5 μm, and sections were mounted on glass slides. Slides were stained with hematoxylin and eosin (H&E) using standard procedures. Immunostaining was performed on 5 μm paraffin sections. Slides were baked at 60°C for 20 min, deparaffinized in xylene, and rehydrated through graded ethanol to water. Heat‐induced epitope retrieval was carried out in 10 mM sodium citrate buffer (pH 6.0, 0.05% Tween‐20) at 95°C–98°C using a steamer (IHC world) for 60 min, followed by cooling to room temperature and PBS washes. Sections were permeabilized in PBS containing 0.1% Triton X‐100 for 15 min and blocked for 1 h at room temperature in PBS with 10% normal serum and 3% BSA. Primary antibodies diluted in blocking buffer were applied overnight at 4°C in a humidified chamber. After three PBS washes, species‐appropriate fluorophore‐conjugated secondary antibodies were applied for 1 h at room temperature, followed by DAPI nuclear counterstain. Slides were rinsed, mounted with Prolong Gold antifade mounting media (Invitrogen), cured, and imaged on a fluorescence microscope using identical acquisition settings across genotypes. Primary antibodies were used at the following dilutions: NCC (ab3553, MilliporeSigma), 1:300; THP (sc‐271022, Santa Cruz Biotechnology), 1:150; AQP2 (sc‐515770, Santa Cruz Biotechnology), 1:150; Ki‐67 (550609, BD Biosciences), 1:400; FITC‐LTA (FL‐1321‐2, Vector Laboratories), 1:1000 and LEA (L32480, Thermo Fisher Scientific), 1:2500. Secondary antibodies conjugated with Alexa fluor (A‐11004 or A‐11036, Thermo Fisher Scientific) were used in the corresponding combinations at dilutions of 1:2500–1:5000. Fluorescence images were acquired at 20× using a Leica DM5000 microscope (Leica Microsystems). Images were initially captured in grayscale and subsequently processed in Adobe Photoshop to perform pseudocoloring and adjust image intensities. Experiments were performed in three biological replicates. Controls were littermate controls that did not carry a homozygous deletion of Bicc1. These included animals with the following genotypes: Cdh16‐cre, Bicc1 ^E4fl/+ , Bicc1 ^E4fl/+ ; Cdh16‐cre, Bicc1 ^E45fl/+ , or Bicc1 ^E45fl/+ ; Cdh16‐cre.

Disclosure

Dr. W. H. Wilson Tang has served as a consultant for Cardiol Therapeutics, Zehna Therapeutics, WhiteSwell, CardiaTec Biosciences, Alleviant Medical, Salubris Biotherapeutics, BioCardia, Tenax Therapeutics, BridgeBio Pharma, Vasa Therapeutics and has received honoraria from Springer and Belvoir Media Group. All other authors have no relationships to disclose.

Liu, C.‐F. , Leon S., Herrig I., Wessely O., and Tang W. H. W.. 2026. “Generation of Mice Harboring Bicc1 Conditional Null Alleles.” genesis 64, no. 1: e70038. 10.1002/dvg.70038.

Data Availability Statement

Data sharing not applicable to this article as no datasets were generated or analysed during the current study.