IL1R2 Marks and Sorts a Spermatogonial Stem Cell Population Critical for Restoring Spermatogenesis Upon Interruption in Mammals
Pengyu Li, Zexuan Zhang, Zihan Xie, Cheng Jin, Zhipeng Wang, Changwei Yuan, Jun Zhu, Jielin Tang, Mengzhen Li, Dingfeng Zou, Xinyu Mang, Jun Liu, Dingyao Chen, Qi Geng, Yan Lu, Ning Zhang, Shiying Miao, Jing Peng, Kai Li, Wei Song
TL;DR
This study identifies IL1R2 as a marker for spermatogonial stem cells that can restore sperm production after disruption, offering a potential therapy for infertility.
Contribution
The discovery of IL1R2 as a specific marker for functionally active spermatogonial stem cells in both mice and humans.
Findings
IL1R2 marks a subpopulation of spermatogonial stem cells critical for restoring spermatogenesis.
IL1R2+ stem cells proliferate via the PI3K-AKT-mTORC1 pathway after spermatogenic disruption.
PI3K-AKT-mTORC1 agonists enhance recovery of spermatogenesis following disruption.
Abstract
Self‐renewal and differentiation of spermatogonial stem cells (SSCs) are critical for sustaining spermatogenesis in adult mammals. However, SSCs are highly heterogeneous, comprising a complex array of subpopulations whose identities and dynamic transitions remain greatly underappreciated. Through in silico analysis, we identified IL1R2 as a surface marker specific to the SSC subpopulation. IL1R2 enables the specific sorting of functionally active SSCs in both human and mouse. Il1r2 CreERT2/+ Rosa26 mTmG/+ mice allowed us to pulse‐label and trace the lineage of Il1r2‐expressing cells. We confirmed that IL1R2+ SSCs support spermatogenesis via both self‐renewal and differentiation. Following spermatogenic disruption, IL1R2+ SSCs are reactivated for proliferation via the PI3K‐AKT‐mTORC1 pathway to replenish the SSC pool. Importantly, we demonstrated that PI3K‐AKT‐mTORC1 agonists can…
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FIGURE 7- —National Key Research and Development Program of China10.13039/501100012166
- —Chinese Academy of Medical Sciences (CAMS) Innovation Fund for Medical Sciences
- —National Natural Science Foundation of China10.13039/501100001809
- —State Key Laboratory Special Fund
- —Beijing Research‐based Ward Excellence Program in Clinical Research
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Taxonomy
TopicsSperm and Testicular Function · Reproductive Biology and Fertility · Pluripotent Stem Cells Research
Introduction
1
Spermatogonial stem cells (SSCs) drive continuous spermatogenesis under homeostatic conditions and mediate fertility restoration following injury in adult mammalian testes [1, 2, 3]. SSCs exhibit remarkable heterogeneity and dynamic regulatory properties, with a series of marker discoveries providing critical insights into their regulation [4, 5]. A subset of A_single_, marked by ID4, PAX7, FOXC2, and PDX1, possesses long‐term self‐renewal capacity [4, 6, 7, 8, 9]. Lineage tracing studies further confirm that GFRα1^+^、EOMES^+^、T^+^、NANOS2^+^、PLVAP^+^、SHISA6^+,^ and BMI1^+^ subsets represent highly potent SSCs with robust self‐renewal capacity and full differentiation potential [10, 11, 12, 13, 14].
Recent advances in single‐cell RNA sequencing (scRNA‐seq) of adult mouse and human testes have underscored the critical role of SSCs in male fertility restoration. Specific subsets, including ID4^+^, PAX7^+^, and FOXC2^+^ SSCs, enter a proliferative state essential for recovery of spermatogenesis [7, 8, 10, 15]. In addition to the expansion of surviving SSCs, transit‐amplifying progenitors such as NGN3^+^ and MIWI2^+^ cells appear capable of dedifferentiation and replenishment of the SSC pool, although their regenerative capacity is lower than that of the aforementioned SSC subsets [16, 17]. While these studies have advanced our understanding of the dynamic SSC changes involved in regeneration, the precise cellular pathways underlying SSC regenerative responses remain poorly characterized. Following chemotherapy‐induced damage, surviving A_undiff_ initiate regenerative responses by activating key signaling pathways, including PI3K/AKT‐mediated FOXO1 inactivation and mTORC1 activation [18]. However, the specific SSC subpopulations that respond to regeneration cues and how these responses are molecularly regulated remain unclear.
Surface markers offer unique advantages for probing SSC biology due to their accessibility for live‐cell interrogation or functional enrichment of intact populations, though only a small number have been characterized in detail. Classic examples include GFRα1, a receptor activated by GDNF, which is crucial for SSC homeostasis — lineage tracing and transplantation highlighting its dynamic behavior [19]; TSPAN8, whose expression distinguishes SSCs with regenerative and differentiation potential, characterized by distinct epigenetic and transcriptional states [9]; THY1 (CD90) enriches for the A_undiff_ population and improves colonization efficiency, albeit with limited specificity as it also captures non‐stem cells [8, 20]; Despite these advances, a clear dissection of SSC heterogeneity and translational potential of these cells remains limited, underscoring the need for additional surface markers to achieve high‐purity isolation, detailed characterization, and precise lineage tracing of SSC subpopulations.
In this study, we identified IL1R2, a previously uncharacterized surface receptor that serves as a conserved and functionally unique marker of SSCs in both mouse and human testes. Compared to broad markers, IL1R2 defines a smaller, more precise subset of SSCs that are highly enriched for functional activity. Mechanistically, we found that PI3K‐AKT‐mTORC1 activation is required for IL1R2^+^ SSC‐mediated regeneration and contributes to replenishing the GFRα1^+^ SSC pool. Our findings provide a unique tool for dissecting SSC heterogeneity and reveal a key transitional subpopulation that links quiescent stem cells and activated progenitor cells during regeneration.
Results
2
Il1r2 Expression Marks a Population of Adult Undifferentiated Spermatogonia
2.1
To identify robust surface markers for SSCs, we analyzed scRNA‐seq data from testicular samples of humans, mice, and busulfan‐induced regeneration models (Figure 1A). In adult human testes, unbiased clustering and UMAP projection revealed a continuous spectrum of germ cell differentiation alongside various somatic populations (Figure S1A). Based on canonical marker expression, testicular cells were categorized into distinct germ cell stages, including undifferentiated spermatogonia (A_undiff_), differentiating spermatogonia, spermatocytes, and late spermatids, as well as somatic cell types like Sertoli cells, Leydig cells (Figure S1B) [21]. Re‐clustering of A_undiff_ identified three transcriptionally distinct subclusters (Figure S1C,D), with cluster 0 corresponding to early‐stage SSCs enriched for stemness‐associated genes such as BCL6, UTF1, FGFR3, GFRα1, and ID4 (Figure S1E,F). Highly variable genes specific to cluster 0 were selected as candidate gene set 1 for surface marker screening (Figure S1G). To assess cross‐species conservation, we analyzed single‐cell transcriptomic data from THY1^+^ A_undiff_ isolated from adult mouse testes. UMAP identified five spermatogonial clusters (Figure S1H), with cluster 0 corresponding to primitive SSCs marked by Foxc2, Gfrα1, Plvap, Id4, Eomes, and Pdx1 (Figure S1I). Trajectory analysis confirmed the gradual transition from SSCs to A_undiff_ (Figure S1J,K). Highly variable genes enriched in cluster 0 were defined as candidate gene set 2 for marker identification (Figure S1L).
Il1r2 is expressed in undifferentiated spermatogonia. (A) Schematic illustration of the technical workflow used to identify and characterize IL1R2+ cells in homeostatic and regenerative spermatogenesis. (B) A Venn diagram reveals the intersection between the five data sets to identify common differentially expressed genes (DEGs). (C) Gene expression heatmap of SSC candidate markers (including IL1R2, LDLR, CD9, APLP2, GFRα1, and RAMP2) in different SSC subtypes. (D) Whole‐mount images of a part of adult mouse (P56) seminiferous tubules showing IL1R2 (green) signals and the IL1R2+ cell distribution illustrated below. Scale bar: 50 µm. (E) Quantification of IL1R2+ cells within different spermatogonial units, demonstrating their predominant presence in As and Apr populations. (F) Immunofluorescence (IF) staining for ZBTB16, FOXC2, LIN28A, WT1 (red), IL1R2 (green), and DAPI (blue) of testis sections from wild‐type adult mice (P56). Scale bar: 50 µm. (G) Immunofluorescence (IF) staining for ZBTB16 (green), SSEA4 (purple), and IL1R2 (gray) of seminiferous tubules from adult humans. Scale bar: 50 µm. (H) Representative flow cytometry gating strategy for isolating subpopulations of undifferentiated spermatogonia (Aundiff). (I) qPCR analysis of Gfrα1, Foxc2, and Il1r2 mRNA expression in sorted populations a, b, c, and an unsorted (u) fraction. Data represent mean ± s.e.m. from n = 3 biological replicates. Student's t‐test; *** p < 0.001.
To explore the plasticity of SSCs under stress, we analyzed A_undiff_ from busulfan‐treated and control mice. UMAP integration under steady‐state and regenerative conditions revealed three SSC subpopulations: SSCs, transitional SSCs, and progenitors (Figure S2A). Regeneration induced changes in SSC composition, with an increase in transitional SSCs and a decrease in SSCs (Figure S2B). Foxc2, Gfrα1, Plvap, and Id4 were enriched in the SSC compartment (Figure S2C,D), accompanied by delayed differentiation (Figure S2E,F). Notably, core markers Foxc2, Plvap, Eomes, and Gfrα1 remained enriched in regenerative SSCs (Figure S2G,H). Differentially upregulated genes in regenerative SSCs were designated as candidate gene set 3 (Figure S2I,J). In addition, receptor databases from mouse and human SSCs provided two more candidate sets (sets 4 and 5) [22]. By intersecting all five sets, we identified six common candidate genes—Ldlr, Il1r2, Cd9, Aplp2, Gfrα1, and Pamp2 (Figure 1B,C; Table S2). Established core markers, including GFRα1 and CD9, were enriched in the SSC population, and through analysis of mouse and human SSC gene sets under various conditions, we found IL1R2 to be a highly conserved marker with significant potential for selectively enriching functionally active SSCs, not previously reported in reproductive studies.
In mouse testes, whole‐mount immunofluorescence showed that IL1R2⁺cells exhibited either single or syncytial staining patterns along the seminiferous tubules (Figure 1D). Based on morphology, most of the IL1R2^+^ spermatogonia were A_s_ (40.3%), 31.3% were A_pr_, and longer syncytia were rare, including A_al‐4_ and A_al‐8_ (Figure 1E). IL1R2 exhibited co‐localization with other stem/ progenitor cell markers (Figure 1F). Quantitative analysis revealed that among IL1R2^+^ cells, 63.8% co‐expressed ZBTB16, 51.7% LIN28A, and 22.4% FOXC2 (Figure S3A), while IL1R2^+^ cells accounted for 18.9% of ZBTB16^+^, 35.9% of LIN28A^+,^ and 26.1% of FOXC2^+^ populations (Figure S3B). A subset of IL1R2^+^ cells exhibited negative staining for MKI67 (Figure S3C). These findings indicate that IL1R2 labels a partially overlapping but transcriptionally distinct SSC subset with a distribution pattern differing from classical stage markers. In human seminiferous tubules, IL1R2 also co‐localizes to varying degrees with both SSEA4^+^ (a marker for human SSCs) and ZBTB16^+^ (a marker for SSCs and A_undiff_) populations (Figure 1G). These results suggest that the expression of Il1r2 in the seminiferous tubule was limited to A_undiff_. Comparing the different levels of gene expression in the cells of the flow circle, we developed a strategy for fluorescence‐activated cell sorting (FACS), which expanded the IL1R2^+^ population horizontally based on E‐CADHERIN and C‐KIT, whose products are on the cell surface (Figure S3D–G). Specifically, the IL1R2^+^ A_undiff_ subpopulation, noted as subgroup a, was exclusively E‐CADHERIN^+^, with significant upregulation in the expression of genes involved in the self‐renewal of stem cells, including Eomes, Id4, Foxc2, and Gfrα1. In contrast, progenitor markers, such as Ngn3, Sox3, and Rary, marked the IL1R2^−^ population as subgroup b, whereas the C‐KIT^+^ subpopulation (subgroup c) was distinguished by Stra8, c‐Kit, Sohlh1, and Sohlh2, whose dynamic expression during spermatogenesis was important for the transition from stemness to differentiation (Figure 1H,I; Figure S3H). To further validate IL1R2 as a surface marker of A_undiff_, we performed flow cytometry analysis on testes from postnatal day 7 (P7) mice, followed by immunofluorescence staining of sorted cells (Figure S3I–M). Subsequent qPCR analysis revealed that IL1R2^+^ cells within the E‐CADHERIN^+^ population exhibited significantly higher expression of SSC‐associated genes, while the expression of differentiation markers was markedly lower (Figure S3N). These results confirm that IL1R2^+^ cells were transcriptionally aligned with early A_undiff_.
IL1R2+ Aundiff Population Supports Spermatogenesis in Adult Mice
2.2
We generated a mouse strain for lineage tracing of IL1R2^+^ cells. The Il1r2 ^CreERT2/+^; *Rosa26^mTmG/+^
- strain was created by crossing Il1r2 ^CreERT2^ mice with Rosa26 ^mTmG^ mice for genetic lineage tracing, where 4OH‐tamoxifen (4OH‐TAM) was administered to induce sustained membrane‐bound GFP expression in specifically labeled IL1R2^+^ cells (Figure 2A; Figure S4A). To evaluate the genetic model, we performed PCR and q‐PCR analyses. PCR validation confirmed the presence of both wild‐type and mutant alleles in Cre‐positive mice (Figure S4B,C). q‐PCR showed no significant difference in mRNA expression between Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ and control mice (Figure S4D), confirming the successful generation of Il1r2 ^CreERT2/+^ mice. Immunofluorescence analysis and flow cytometry revealed that 85.7% of GFP^+^ cells were IL1R2^+^ cells three days after 4OH‐TAM treatment, confirming the accuracy of the genetic lineage tracing model (Figure S4E–G). Most GFP clones were either A_s_ or A_pr_ spermatogonia, with a few long‐chain cells, among which A_s_ spermatogonia had the highest labeling frequency (Figure S4H). Notably, 89.3% of GFP^+^ cells were GFRα1^+^, whereas only 32.1% of GFRα1^+^ cells were GFP^+^ (Figure 2B–D). Additionally, single‐cell data analysis revealed that IL1R2^+^ cells make up 27.27% of the GFRα1^+^ cell population (Figure 2E,F). Moreover, IL1R2 was co‐expressed with other well‐established SSC markers, including EOMES and PAX7 (Figure S4I). Despite the substantial overlap, the GFP/ IL1R2 signal could be clearly distinguished from those of other SSC markers like ZBTB16 and GFRα1 (Figure 2B). Therefore, IL1R2^+^ cells are likely to represent a distinct subpopulation within the A_undiff_ group.
*IL1R2+SSC can maintain spermatogenesis. (A) Schematic of the lineage tracing experiment in Il1r2 CreERT2/+; Rosa26 mTmG/+ mice. (B) Immunofluorescence (IF) staining for ZBTB16 (gray), GFRα1 (red), and GFP (green) of testis sections from Il1r2 CreERT2/+; Rosa26 mTmG/+ mice. White arrows indicate ZBTB16+/ GFRα1+/ GFP+ cells, while red arrows indicate ZBTB16+/ GFRα1+/ GFP− cells. Insets show higher magnifications of the boxed areas on the right. Scale bar: 50 µm. (C) Quantification of GFRα1+ and GFRα1−cell fractions in the GFP+ cell at Day 3 post‐induction. n > 50 tubules each from five independent males. (D) The proportion of GFP+ cells in the GFRα1+ population at Day 3 post‐induction. n > 50 tubules each from five independent males. (E) UMAP representation showing the distribution of Il1r2
- cells (red) and Gfrα1
- cells (green) in the dataset. (F) Bar plot showing the fraction of Gfrα1
- cells that are either Il1r2
- or Il1r2 − (blue and light blue bars, respectively). (G) Schematic illustration of the lineage tracing workflow for IL1R2+ cells. Experimental timeline indicating time points (D7, D28, D56, D120, and D365) for sample collection post‐4OH‐TAM induction. (H) Immunostaining for DAPI (blue) and ZBTB16 (purple) showing the progression of GFP+ cells (green) in seminiferous tubules at days 7 (D7), 28 (D28), and 56 (D56) after 4OH‐TAM induction. The left image of the last row shows GFP‐positive seminiferous tubules filled with mature sperm. The middle image displays GFP‐positive spermatozoa isolated from the epididymis. The right image shows the GFP‐positive offspring. Scale bars: 50 µm. (I) Bar graph showing the proportion of GFP+ and GFP− offspring from Il1r2 CreERT2/+; Rosa26 mTmG/+ lineage tracing mice (n = 5) following tamoxifen induction and natural mating. (J) The testes (scale bar: 1 mm) and seminiferous tubules (scale bar: 50 µm) observed on days 120 (D120) and 365 (D365) post‐TAM induction. (K, L) Proportion dynamics of clone size (K) and GFP patches (L). Values represent mean ± s.e.m. n > 30 tubules each from five independent males.*
Further lineage tracing experiments allowed us to track the progression of spermatogenesis in Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice on days 7, 28, 56, 120, and 365 after 4OH‐TAM treatment (Figure 2G). A_s_ coexisted with both A_pr_ and A_al_ spermatogonia (interconnected cells in syncytial cysts 4 and 8) on day 7 and day 28. The GFP signal was more enriched in the syncytial cysts consisting of both ZBTB16^+^ and ZBTB16^−^ populations. Morphologically, the progeny of the labeled germ cells appeared as differentiated spermatogonial clusters. On day 56, GFP^+^ sperm were detectable, which gave rise to GFP‐fluorescent offspring, confirming that IL1R2^+^ SSCs can produce all the progeny to support successful spermatogenesis (Figure 2H,I). As revealed in the extended lineage‐tracing experiment up to days 120 and 365, the GFP^+^ cells developed as a continuous and persistent plaque (Figure 2J). Over time, the number of GFP clones decreased, but the average clone length increased across all stages of differentiation in some lumens, further supporting the notion that IL1R2^+^ SSCs contribute to continuous spermatogenesis in adult mammals (Figure 2K,L).
Ablation of IL1R2+ SSCs Impairs Spermatogenesis and Long‐Term Fertility in Mice
2.3
To evaluate the physiological needs of IL1R2^+^ SSC during spermatogenesis, we constructed the Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ strain in which IL1R2^+^ cells expressed diphtheria toxin (DTA) upon 4OH‐TAM administration (Figure 3A). On day 56 post‐4OH‐TAM treatment, the percentage of IL1R2^+^ cells decreased to 0.018% of total testicular cells, whereas IL1R2^+^ cells accounted for 0.047% of total cells in the control group, thus confirming the success of ablation (Figure S5A,B). To assess the functional role of IL1R2^+^ cells in long‐term fertility, we conducted breeding assays. In 4OH‐TAM‐treated Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ males, offspring numbers were significantly reduced, whereas all control males showed normal fertility (Figure S5C).
Ablation of IL1R2+ SSC in mice results in a disruption of spermatogenesis. (A) 4OH‐TAM activates CreERT2 in Il1r2 CreERT2/+; Rosa26 DTA/+ mice, inducing DTA expression and ablation of IL1R2+cells (Top). 4OH‐TAM or vehicle (DMSO) was administered at postnatal day 56 (P56), and samples were collected 56 days later (D56) to ensure complete clearance of differentiated germ cells derived from IL1R2+ SSC (Bottom). (B) In adult mice, the testes of Il1r2 CreERT2/+; Rosa26 DTA/+ mice were smaller than those of Rosa26 DTA mice. (C) Assessment of testis to body weight ratio (mg/g) in Rosa26 DTA and Il1r2 CreERT2/+; Rosa26 DTA/+ adult mice; n = 6, Student's t‐test; ∗∗∗∗ p < 0.0001. (D) Hematoxylin and eosin (HE) staining of paraffin‐embedded sections from the testis and epididymis of Rosa26 DTA and Il1r2 CreERT2/+; Rosa26 DTA/+mice, * indicates degenerated tubules. Scale bar: 50 µm. (E) Assessment of seminiferous tubule area from Rosa26 DTA and Il1r2 CreERT2/+; Rosa26 DTA/+ adult mice; n > 20 tubules each from three independent males, Student's t‐test; **** p < 0.0001. (F) Assessment of sperm number from Rosa26 DTA and Il1r2 CreERT2/+; Rosa26 DTA/+ adult mice; n = 3 animals, Student t‐test; ∗∗∗ p < 0.001. (G) Immunofluorescence (IF) staining for GFRα1, C‐KIT (red), and ZBTB16, WT1 (green) of testis sections from Il1r2 CreERT2/+; Rosa26 DTA/+ mice and Rosa26 DTA mice. Scale bar: 50 µm. (H–K) Statistical analysis of the number of GFRα1+ cells (H), ZBTB16+ cells (I), C‐KIT+ cells (J), and WT1+ cells (K) per tubule in Il1r2 CreERT2/+ Rosa26 DTA/+ mice and Rosa26 DTA mice; n > 50 tubules from each of three independent males; Student's t‐test; ∗∗ p < 0.01; **** p < 0.0001; ns, no significance.
These results confirm that ablation of IL1R2^+^ cells compromises the ability to maintain continuous spermatogenesis, ultimately leading to progressive infertility. Compared to the control group, testicular volume and weight were significantly reduced in Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ mice (Figure 3B,C). Histological analysis revealed that spermatogenesis remained unaltered in the control group upon 4OH‐TAM exposure, whereas nearly all cavities appeared vacuolized, with the cross‐sectional area of seminiferous tubules clearly reduced in Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ mice, indicating that spermatogenesis was severely disrupted owing to the loss of the IL1R2^+^ population (Figure 3D,E). Furthermore, the number of sperm in a few cavities of the epididymis was the least affected in Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ mice, accounting for only 40.1% of the control (Figure 3D,F). In these cavities, GFRα1^+^ SSCs, ZBTB16^+^ A_undiff_ and C‐KIT^+^ A_diff_, though greatly reduced in number, remained detectable at the basement membrane of seminiferous tubules (Figure 3G–J). Consistently, a clear reduction was observed in the number of spermatocytes (SYCP3^+^), round sperm cells (PNA^+^), and germ cells (TRA98^+^) in Il1r2 ^CreERT2/+^; Rosa26 ^DTA/+^ mice (Figure S5D–G), whereas Sertoli cells remained unaffected (Figure 3K). Taken together, we determined that the IL1R2^+^ SSC population is critical for spermatogenesis in adult mice.
Development of IL1R2+ SSCs Under Physiological Conditions
2.4
We studied the behavior of the IL1R2^+^ SSCs population across the entire span of spermatogenesis. From days 2 to 20 post‐4OH‐TAM induction, the seminiferous tubules were stained for GFRα1, SOX3, and GFP, and the number of GFP^+^ cells was quantified (Figure 4A). Shortly after induction (D2), all the labeled clones contained one or two GFRα1^+^ cells; up to D14, GFP^+^/ GFRα1^−^ cells emerged. At D20, most GFP^+^ cells exhibited positive staining for C‐KIT but negative for GFRα1, suggesting that the GFP^+^/ GFRα1^+^ population differentiated into the GFRα1^−^ progeny over this period of time (Figure 4B). This showed a highly variable regular pattern of SSC development. The dynamic transition was particularly evident from D2 to D6 (Figure 4C). Since SOX3 has been linked to the differentiation of GFRα1^+^ SSCs, we cross‐checked the dynamics of GFP^+^, GFRα1^+^, and SOX3^+^ populations (Figure 4D). The overall SSC population appeared to be highly heterogeneous, primarily comprising of GFRα1^+^/ GFP^−^/ SOX3^−^ (No. 1), GFRα1^+^/ GFP^+^/ SOX3^−^ (No. 2); GFRα1^+^/ GFP^+^/ SOX3^+^ (No. 3), and GFRα1^−^/ GFP^−^/ SOX3^+^ (No. 4) subpopulations. This was further supported by the quantification of GFP^+^ cells (Figure 4E). After 4OH‐TAM induction, the GFRα1^+^/ GFP^+^/ SOX3^−^ population, likely representing labeled, self‐renewing SSCs, predominated at D2. Meanwhile, the GFRα1^+^/ GFP^+^/ SOX3^+^ population, indicating SSCs reactivated for differentiation, began to emerge. From D2 to D4, the self‐renewing population decreased, while the differentiating population steadily increased. From D4 onward, two new populations emerged and rapidly increased in size, both of which likely comprised differentiating SSCs, with more GFRα1^−^/ GFP^+^/ SOX3^+^ cells during the early stages and more GFRα1^−^/ GFP^+^/ SOX3^−^ cells at subsequent stages. In contrast, GFRα1^+^/ GFP^+^/ SOX3^−^ cells maintained a slow decline slowly followed by GFRα1^+^/ GFP^+^/ SOX3^+^ cells from D6 when the population peaked. From D6 onward, GFRα1^−^/ GFP^+^/ SOX3^+^ and GFRα1^−^/ GFP^+^/ SOX3^−^ became the dominant populations with the latter comprising the largest population from D10. Therefore, during homeostasis, cell types 1, 2, 3, and 4 constituted a functional hierarchy arranged in this order. The heterogeneity was most likely the result of both pulsed‐chase labeling and spermatogenesis. The changes were also reflected in the total increase in the number of GFP^+^ cells over the course of the experiment (Figure 4F), as well as the increase in GFRα1^−^ cells while the number of GFRα1^+^ cells did not change (Figure 4G,H). These findings confirmed that GFP^+^ (IL1R2^+^) SSCs expanded and differentiated into GFP^+^ progenies, contributing to the maintenance of spermatogenesis during homeostasis. Consistently, C‐KIT, a marker for differentiation, increased the number of cells remarkably throughout the experiment, particularly from D6 (Figure 4I) [23]. Furthermore, the overall trend was in line with the dynamics of the cells at each individual stage of spermatogenesis (Figure 4J). Typically, GFP^+^ A_s_, initially the dominant group, declined over time and then persisted in relatively low proportions, in contrast to GFP^+^ A_al_. Overall, IL1R2^+^ SSCs were able to complete the transition from self‐renewal to differentiation and further differentiate into all progeny, which is critical for maintaining the homeostasis of SSCs and supporting continuous spermatogenesis in adult mice.
Developmental fate analysis of IL1R2+SSC during homeostasis. (A) Schematic illustration of the lineage tracing workflow for IL1R2+ cells. (B) Immunofluorescence (IF) staining for GFRα1 (red) and GFP (green) of a wholemount from Il1r2 CreERT2/+ Rosa26 mTmG/+ mice. Scale bar: 50 µm. (C) Clonal dynamics of average clone composition labeled by GFRα1 and C‐KIT following induction in GFP+ population. n > 50 tubules from each of the three independent males. (D) Immunofluorescence (IF) staining for GFRα1 (purple), SOX3 (red), and GFP (green) of a whole mount from Il1r2 CreERT2/+; Rosa26 mTmG/+ mice. Scale bar: 50 µm. (E) Clonal dynamics of average clone composition labeled by GFRα1 and SOX3 following induction in GFP+ population. n > 50 tubules from each of three independent males. (F) Distribution of clone size from days 2 to 20 after induction, scored by the number of GFP+ cells. (G–I) Fraction of GFRα1+ cells (G), GFRα1−cells (H), and C‐KIT+ cells (I) out of the total GFP+ population from days 2 to 20 after induction. n > 50 tubules from each of the three independent males. (J) Kinetics of the overall composition of pulse‐labeled GFP+ cells in different Aundiff subtypes. n > 50 tubules from each of the three independent males.
IL1R2+ SSCs Restore Spermatogenesis upon Interruption
2.5
Next, we assessed the capability of IL1R2^+^ SSCs to regenerate spermatogenesis in a busulfan‐induced testicular injury model (Figure 5A) [24]. ScRNA‐seq analysis revealed an increased expression of Il1r2 in the SSC subpopulation of the regeneration group following busulfan treatment (Figure S6A,B). FACS analysis showed that the size of C‐KIT^+^ A_diff_ cells decreased significantly after busulfan treatment, indicating the exhaustion of spermatogonia. The IL1R2^+^ SSCs appeared resistant and responsive to busulfan, since their size increased from 0.042% to 0.083% (Figure S6C). We conducted a comprehensive analysis to elucidate the fate of IL1R2^+^ SSC populations during the reestablishment of spermatogenesis following busulfan treatment. First, we administered various concentrations of busulfan to wild‐type mice and observed changes in the repair kinetics of IL1R2^+^ SSCs. We observed a progressive transition of IL1R2^+^ SSCs from a quiescent to a proliferative state with increasing busulfan concentrations (10, 20, 30, and 40 mg/kg). Correspondingly, the percentage of MKI67^+^ cells within the IL1R2^+^ SSC population increased, reaching 80.32%, 96.45%, and 100% at the respective doses (Figure S6D,E). Subsequently, eight‐week‐old Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice were pulse‐labeled with 4OH‐TAM and treated with 10 mg/kg busulfan to induce spermatogenic regeneration. By day 20 post‐treatment, some GFP^+^ cells formed small regenerative clusters and all IL1R2^+^ SSCs appeared to be committed to spermatogenic recovery (Figure 5B). Following busulfan exposure, we observed a significant increase in the proportion of IL1R2^+^ SSCs entering the cell proliferation phase during the recovery phase compared to that in the homeostatic phase (Figure 5C). These findings suggested that IL1R2^+^ SSCs are capable of responding to the toxic effects of chemical agents and initiating spermatogenic restoration by increasing the number of proliferating cells. After busulfan treatment, the number of GFP^+^ cells undergoing death progressively increased. By day 20 of treatment, the GFP^+^ cells began to recover; however, the cell count remained significantly different from the baseline homeostatic level. Alterations in these two variables could serve as indicators of the selective elimination and attenuation of IL1R2^+^ cyst formation (Figure 5D,E). Additionally, our data revealed a biphasic pattern of cell dynamics within the GFRα1^+^ population, characterized by an initial decline followed by an increase in the proportion of GFP^+^ cells. During the recovery phase, a significant increment in the number of GFRα1^+^ cells was observed compared to the homeostatic state (Figure 5F). Throughout the process of homeostatic spermatogenesis, IL1R2^+^/ GFRα1^+^ cells maintained a consistent presence. In the regenerative phase, the percentage of GFRα1^+^ cells within the GFP^+^ population was elevated compared to that observed during homeostatic spermatogenesis, which indicated that most of the IL1R2^+^ SSCs that persisted following busulfan treatment were GFRα1‐positive. Moreover, we found a substantial increase in the proportion of GFRα1^+^ A_s_ spermatogonia that engaged in the regenerative process within the GFP^+^ population (Figure 5G). Our findings suggested that IL1R2^+^ SSCs can self‐renew and proliferate during the recovery phase following injury. This capacity enables them to replenish the GFRα1^+^ SSC pool, thereby significantly contributing to the regenerative potential of the stem cell bank. To assess the long‐term efficacy of IL1R2^+^ SSCs in promoting spermatogenic recovery, we evaluated spermatogenic recovery for up to 60 days after busulfan treatment (Figure 5H). The analysis revealed an increase in the number of colonies in pulse‐labeled Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice treated with busulfan, albeit with a significantly reduced colony size. These findings suggested a persistent effect of IL1R2^+^ SSCs on spermatogenic regeneration following busulfan‐induced injury (Figure 5I–K). Collectively, the data suggested that IL1R2^+^ SSCs contribute significantly to the restoration of spermatogenesis following busulfan‐induced damage.
Developmental fate analysis of IL1R2+SSC during injury. (A) Schematic illustration for the lineage tracing workflow of Il1r2 CreERT2/+; Rosa26 mTmG/+ mice following short‐term induction using 4OH‐TAM and Busulfan. (B) Immunofluorescence (IF) staining for GFP (green) and MKI67 (yellow) of a whole mount from Il1r2 CreERT2/+; Rosa26 mTmG/+ mice after Busulfan treatment. Scale bar: 50 µm. (C) The proportion of MKI67+ cells in the GFP+ population in both the homeostatic and regenerative states. n > 50 tubules from each of the three independent males; Student t‐test; ∗∗ p < 0.01; ∗∗∗ p < 0.001. (D) Immunofluorescence (IF) staining for GFP (green), GFRα1 (red), ZBTB16 (yellow), and DAPI (blue) of the whole‐mount from Il1r2 CreERT2/+; Rosa26 mTmG/+ mice after Busulfan treatment. Scale bar: 50 µm. (E) Number of GFP+ cells in Il1r2 CreERT2/+ Rosa26 mTmG/+ mice from days 0 to 20 post‐Busulfan induction. n > 50 tubules from each of the three independent males; Student's t‐test; ∗∗∗ p < 0.001; ns, no significance. (F, G) The proportion of GFRα1+ cells (F) and GFRα1+ As cells (G) in the GFP+ population in both the homeostatic and regenerative states from days 0 to 20 post‐Busulfan induction. n > 50 tubules from each of the three independent males; Student's t‐test; ∗ p < 0.05; ∗∗ p < 0.01; ∗∗∗ p < 0.001; ns, no significance. (H) Schematic illustration of the lineage‐tracing workflow for Il1r2 CreERT2/+ Rosa26 mTmG/+ mice following long‐term induction with 4OH‐TAM and Busulfan. (I) Testes (scale bar: 1 mm) and seminiferous tubules (scale bar: 50 µm) of 4OH‐TAM‐stimulated Il1r2 CreERT2/+; Rosa26 mTmG/+ mice on day 60 post‐Busulfan induction. (J, K) Proportion dynamics of clone size (J) and GFP+ patches (K). Values represent mean ± s.e.m. n > 20 tubules from each of the three independent males; Student t‐test; ∗ p < 0.05; ∗∗∗ p < 0.001.
Activation of the PI3K‐AKT‐mTORC1 Pathway Promotes the Recovery Supported by IL1R2+ SSCs
2.6
To elucidate the repair mechanisms of IL1R2^+^ SSCs following busulfan‐induced injury to spermatogenic cells, we isolated and characterized IL1R2^+^ SSCs from busulfan‐treated and control groups using scRNA‐seq (Figure S6F). We found a significant enrichment of the proliferation marker MKI67, as well as the stem cell markers GFRα1 and FOXC2, in the regenerative subgroup (Figure S6G). DEGs analysis comparing homeostatic and regenerative IL1R2^+^ SSC subsets highlighted the remarkable upregulation of genes associated with cell proliferation within the regenerative subset (Figure S6H). Furthermore, a gene ontology (GO) analysis showed significant enrichment of biological processes related to mitosis and cell cycle among the genes that were highly expressed in the regenerative subgroup (Figure S6I). Gene set enrichment analysis (GSEA) revealed enrichment of the PI3K‐AKT‐mTOR signaling pathway, suggesting that regenerative IL1R2^+^ SSCs may regulate stem cell proliferation through this pathway (Figure 6A). Next, we investigated the role of the PI3K‐AKT‐mTOR pathway in IL1R2^+^ SSC regeneration. Since the PI3K‐AKT‐mTOR pathway is mainly regulated by GDNF, we coated the microbeads with GDNF and Dil and injected them into the testicular interstitium via the rete testis [25, 26]. Immunofluorescence analysis revealed activation of the pathway in IL1R2^+^ SSCs upon GDNF stimulation, as evidenced by the phosphorylation of the downstream protein S6 (P‐S6) (Figure 6B,C). Subsequently, we treated wild‐type mice with busulfan and performed immunofluorescence analysis a few days later (Figure 6D). The results revealed a reduction in the number of nuclear localizations of FOXO1, concurrent with an increase in its cytoplasmic localization within the IL1R2^+^ SSCs (Figure 6E,F). The alterations in FOXO1 localization within the IL1R2^+^ SSCs following busulfan treatment were attributed to activation of the PI3K‐AKT pathway, which hindered FOXO1 nuclear translocation and subsequent functional engagement. Four days after busulfan treatment, the mice were injected with RAPA, an inhibitor of the mTORC1 pathway, to assess its impact on cellular processes (Figure 6D) [27]. Compared to the homeostatic developmental stage, busulfan‐induced damage led to a significant increase in S6 protein phosphorylation. However, RAPA treatment resulted in a substantial reduction in S6 phosphorylation and a notable decrease in the number of IL1R2^+^ SSCs involved in the repair process (Figure 6G–I). Furthermore, DQA, an activator of the PI3K‐AKT‐mTOR pathway, was injected into mice four days after busulfan injection (Figure 6D) [28, 29]. We observed a decrease in the proportion of FOXO1 that exhibited nuclear localization within the IL1R2^+^ SSCs upon DQA treatment, along with an increase in S6 protein phosphorylation (Figure 6J–M). To strengthen the correlation between this signaling axis and IL1R2^+^ SSC regeneration, we performed pharmacological validation in busulfan‐injured mice treated with MHY1485 (mTOR activator) [29, 30] and GDC‐0941 (PI3K inhibitor) [31, 32] (Figure S6J,L). Quantification of P‐S6^+^/ IL1R2^+^ cells and total IL1R2^+^ cells revealed that MHY1485 treatment significantly increased the proportion of P‐S6^+^ cells and promoted the recovery of IL1R2^+^ cells, while GDC‐0941 treatment reduced both parameters (Figure S6K,M,N). Collectively, the results suggested that during the repair of busulfan‐induced injury, IL1R2^+^ SSCs proliferate by activating the PI3K‐AKT‐mTORC1 pathway, which may in turn contribute to the preservation of the stem cell pool.
Role of the PI3K‐AKT‐mTORC1 pathway in IL1R2+SSCs during regeneration. (A) GSEA analysis showing the upregulated pathway activity in IL1R2+ SSC from mouse testes. (B) Immunofluorescence (IF) staining for IL1R2 (gray), P‐S6 (yellow), and Dil (purple) signals of whole mount from mice treated with GDNF or BSA. Scale bar: 50 µm. (C) Fraction of P‐S6+ cells out of the total IL1R2+ population between GDNF‐treated and BSA‐treated groups. n=30 tubules from each of the three independent males; Student t‐test; *** p < 0.001. (D) Schematic illustration for the lineage tracing workflow of WT mice following induction with Busulfan and RAPA, DQA, or DMSO. (E) Whole‐mount images of a part of the seminiferous tubules from WT mice treated with Busulfan or DMSO showing IL1R2 (red) and FOXO1 (green) signals. Scale bar: 50 µm. (F) The intracellular distribution of FOXO1 within IL1R2+ SSC was analyzed from panel E (C, predominantly cytosolic; N, predominantly nuclear; N + C, nuclear and cytosolic). (G) Immunofluorescence (IF) staining for IL1R2 (red) and P‐S6 (green) of the whole‐mount from WT testes after Busulfan or RAPA treatment. Scale bar: 50 µm. (H) The proportion of P‐S6+ cells in the IL1R2+ population in WT mice treated with Busulfan or RAPA. n = 10 tubules from each of the three independent males; Student t‐test; ∗∗∗ p < 0.001. (I) The number of IL1R2+ cells in WT mice treated with Busulfan or RAPA. n = 10 tubules from each of the three independent males; Student t‐test; ∗∗ p < 0.01. (J) Whole‐mount images of a part of the seminiferous tubules from WT mice treated with Busulfan, DMSO, or DQA showing IL1R2 (red) and FOXO1 (green) signals. Scale bar: 50 µm. (K) The intracellular distribution of FOXO1 within IL1R2+ SSC was analyzed from panel J (C, predominantly cytosolic; N, predominantly nuclear; N + C, nuclear and cytosolic). (L) Immunofluorescence (IF) staining for IL1R2 (red) and P‐S6 (green) of whole‐mount from WT testis after Busulfan or DQA treatment. Scale bar: 50 µm. (M) The proportion of P‐S6+ cells in the IL1R2+ population in WT mice treated with Busulfan or DQA. n = 30 tubules from each of the three independent males; Student's t‐test; ∗∗ p < 0.01; ∗∗∗ p < 0.001.
Evolutionary Conservation of the Role of IL1R2+ SSCs
2.7
To assess the functionality of IL1R2^+^ SSCs, we isolated highly enriched IL1R2^+^ and THY1^+^ populations by FACS, which were used for subsequent functional experiments (Figure 7A; Figure S7A). Initially, we found that ZBTB16^+^ A_undiff_ was preferentially enriched following affinity‐sorting with anti‐IL1R2 antibodies (Figure S7B,C). Cells sorted from Rosa26 ^mcherry^ mice were cultured in the SSC‐conditioned medium, and a cluster of clones that emerged was designated SSC colonies (Figure 7B). Cultured under identical conditions, IL1R2^+^ SSCs showed significantly higher colony‐forming efficiency than both unsorted populations and THY1^+^ spermatogonia (Figure 7C). In addition, IL1R2^+^ SSCs, THY1^+^ spermatogonia, and unsorted populations from Rosa26 ^mCherry^ mice were transplanted into busulfan‐treated donor testes, and mCherry^+^ clones were quantified after eight weeks. Despite the presence of mCherry^+^ colonies in the seminiferous tubules derived from all three cell types, IL1R2^+^ SSCs exhibited a significantly higher capacity for clonal formation than unsorted populations and THY1^+^ cells (Figure 7D,E). Thus, IL1R2 serves as a selective marker for enriching SSCs with robust stemness. To characterize IL1R2^+^ SSCs, we isolated IL1R2^+^, THY1^+^, and C‐KIT^+^ cells from mouse testes using FACS for RNA‐Seq analysis. We conducted a comparative analysis of gene expression to assess the differential expression profiles of IL1R2^+^ SSCs, THY1^+,^ and C‐KIT^+^ spermatogonia (Figure S7D). Examination of gene expression profiles revealed that the IL1R2^+^ population exhibited a robust expression of stem cell‐associated genes, including Gfrα1, Eomes, Etv5, and Pdx1. In contrast, genes linked to differentiation processes, such as c‐Kit, Sohlh1/2, Piwil2, Sycp3, and Sycp1, were highly expressed in the THY1^+^ and C‐KIT^+^ populations (Figure 7F). GO analysis was performed on the differentially expressed genes between IL1R2^+^ SSCs and THY1^+^ spermatogonia. The analysis indicated that genes with high expression were significantly enriched for terms related to male gonadal development, seminiferous tubule development, tissue development, and tissue regeneration. Conversely, the downregulated genes were enriched in terms associated with cell cycle and gene transcription processes (Figure S7E). These findings suggested that IL1R2^+^ SSCs are involved in mediating a more crucial effect within the male reproductive system, contributing significantly to development and regeneration, compared to THY1^+^ spermatogonia, which exhibit high proliferation and transcriptional activity. GO analysis was performed on the DEGs between IL1R2^+^ SSCs and C‐KIT^+^ spermatogonia. The upregulated genes were significantly enriched in terms of gamete production, negative regulation of stem cell differentiation, maintenance of stem cell populations, reproduction, and stem cell proliferation. Conversely, the downregulated genes had entries enriched for the meiotic cycle, sperm development, and chromatin remodeling (Figure S7F). These observations indicated that IL1R2^+^ SSCs are primarily responsible for maintaining stemness and play a central role in gametogenesis, surpassing the functions of C‐KIT^+^ spermatogonia. Overall, the results suggested that IL1R2^+^ SSCs possess unique molecular characteristics of stemness and exhibit a high capacity for stem cell function.
IL1R2+SSC exhibited stem cell characteristics in human and mouse testis. (A) Strategy for functional validation of IL1R2+ cells sorted from Rosa tomato mice. (B–E) In vitro SSC culture and transplantation assay. B. Colony formation assays of IL1R2+, THY1+, and unsorted populations. Scale bar =50 µm. C. Statistical analysis of the results presented in (B). D. Examination of donor‐derived spermatogenesis in recipient mice two months after the transplantation, Scale bar =1 mm. E Statistical analysis of the results presented in (D); Student's t‐test; ∗∗ p < 0.01; ∗∗∗ p < 0.001. (F) Heatmap of stemness‐ and differentiation‐associated marker genes in IL1R2+ cells compared with C‐KIT+ cells (top) and THY1+ cells (bottom) from mouse testes. (G) Schematic diagram of the human IL1R2+ cell‐sorting strategy. (H, I) The purity of the sorting was confirmed by a three‐color immunofluorescent assessment for α6‐INTEGRIN, ZBTB16 (red), DDX4 (yellow), and IL1R2 (green) in the sorted cells; Student's t‐test; ∗∗ p < 0.01; ∗∗∗ p < 0.001. (J) Volcano plot of differential gene expression between IL1R2+ SSC versus SSEA4+ SSC. (K) Enriched GO terms in upregulated and downregulated genes in IL1R2+ cells versus SSEA4+ Aundiff. The hypergeometric test was used for statistical analysis. (L) Heatmap of stemness‐ and differentiation‐associated marker genes expressed in IL1R2+ and SSEA4+ cells from the human testes. (M) GSEA analysis showing the upregulated pathway activity in IL1R2+ SSC from human testes.
IL1R2^+^ SSCs were also present in human testes (Figure 1G). We used an antibody against IL1R2 to selectively enrich and isolate these cells from the seminiferous tubules of patients with obstructive azoospermia (OA) by FACS (Figure 7G; Figure S7G). Fluorescence microscopy of the sorted cells confirmed that the IL1R2 antibody facilitated the acquisition of high‐purity human IL1R2^+^ SSCs. Additionally, a partial, but not complete, overlap in expression with ZBTB16 and α6‐INTEGRIN was noted (Figure 7H,I). Subsequently, we performed RNA‐seq and evaluated the differential gene expression profiles of human IL1R2^+^ SSCs, SSEA4^+^ spermatogonia, and C‐KIT^+^ spermatogonia (Figure 7J). GO analysis revealed that genes preferentially upregulated in IL1R2^+^ SSCs were enriched for stem cell proliferation, male gonad development, and stem cell differentiation. Conversely, the downregulated genes were primarily enriched in biological processes associated with cell cycle and oxidative phosphorylation (Figure 7K). Compared to those in C‐KIT^+^ spermatogonia, genes that were downregulated in IL1R2^+^ SSCs were enriched for fertilization, flagellar movement, and cell migration. Conversely, genes upregulated in IL1R2^+^ SSCs were enriched in biological functions, including cell fate determination, growth and development, retinoic acid response, and stem cell proliferation (Figure S7H,I). Heatmap analysis further revealed a relatively higher expression of genes associated with stemness and regeneration in IL1R2^+^ SSCs, whereas genes related to differentiation were expressed at lower levels (Figure 7L; Figure S7J). GSEA of differentially expressed genes between IL1R2^+^ SSCs and SSEA4^+^ spermatogonia highlighted the specific enrichment of the PI3K‐AKT and mTOR signaling pathways, which are involved in stem cell fate (Figure 7M). These findings together suggested that human IL1R2^+^ SSCs exhibit robust molecular characteristics of stem cells and may play a significant role in spermatogenic homeostasis and regeneration.
Discussion
3
Our study establishes IL1R2 as a conserved surface marker, and we reveal the heterogeneity and hierarchical nature of IL1R2^+^ SSCs in the developmental fate of SSCs. IL1R2^+^ cells are capable of both self‐renewal and differentiation. However, as shown in the lineage‐tracing experiments (Figure 2H–J), although long‐term tracking following a single pulse of 4OH‐TAM showed persistent GFP^+^ clones, IL1R2^+^/ GFP^+^ clones gradually declined, indicating that not all IL1R2^+^ SSCs undergo self‐renewal. During homeostasis, a subset of IL1R2^+^ SSCs persistently proliferate and self‐renew, allowing the detection of clonal patches after long‐term pulse labeling. In the declining A_s_ zone, approximately 25% of IL1R2^+^/GFP^+^ A_s_ persist, as observed in short‐term labeling (Figure 4J). Another subset of IL1R2^+^ SSCs differentiates into SOX3^+^ progenitor cells during homeostasis, promoting irreversible differentiation commitment and playing a key role in SSC transplantation and regeneration (Figure 4D). Importantly, IL1R2^+^ SSCs occupy an intermediate stage among developmental phases, consistent with the current model of functional heterogeneity underlying SSC dynamics [21]. The specific loss of IL1R2^+^ SSCs in diphtheria toxin‐mediated ablation leads to a reduction in progeny numbers or infertility. This highlights that IL1R2^+^ SSCs are a functionally indispensable subpopulation of stem cells, crucial for maintaining spermatogenic homeostasis.
Under regenerative conditions, IL1R2^+^ SSCs exhibit markedly increased proliferation and clone formation, similar to other SSC populations such as FOXC2^+^, GFRα1^+^, ID4^+^, EOMES^+,^ and PAX7^+^ cells, indicating their active role in regeneration and stem cell‐mediated repair [7, 8, 10, 15, 19]. The toxicity of busulfan eliminates some subpopulations of A_undiff_, leading not only to a shortening of clone length but also to a reduction in niche crowding [3]. In this process, IL1R2^+^ SSCs shifted from IL1R2^+^/ GFRα1^−^ to IL1R2^+^/ GFRα1^+^, which resulted in the expansion of GFRα1^+^ A_s_ (from 2% to 6%) and enhancing regenerative potential (Figure 5F,G). It is possible that IL1R2^−^/ GFRα1^+^ SSCs differentiate into IL1R2^+^ / GFRα1^+^, or IL1R2^+^ / GFRα1^−^ SSCs turn into IL1R2^+^/ GFRα1^+^ by acquiring proliferative capacity. These findings nevertheless provide key insights into the heterogeneity within GFRα1^+^ SSCs, extend previous lineage‐tracing studies, and are critical for understanding SSCs homeostasis and regeneration [19]. Moreover, the PI3K‐AKT‐mTORC1 pathway is a complex and finely regulated cascade that serves as a crucial regulatory mechanism for stem cell‐driven repair processes in damaged tissues [33, 34]. Our work further demonstrates that the PI3K‐AKT‐mTORC1 pathway is preferentially activated in IL1R2^+^ SSCs during regeneration, underscoring their key role as a driving population in A_undiff_‐mediated repair [18]. Meanwhile, we found that the agonist using the PI3K‐AKT‐mTORC1 pathway can induce IL1R2^+^ SSC expansion to restore depleted SSC pools, underscoring their potential as facilitators of regenerative spermatogenesis, which might become a potential adjunct committed to regenerative spermatogenesis. Taken together, our data strongly suggested that IL1R2^+^ SSCs conferred resistance to chemotherapeutic agents and facilitated endogenous regenerative processes mediated by agonists. However, the underlying mechanism remains elusive. Further understanding of PI3K‐AKT‐mTORC1's role in SSCs regeneration can provide leverage for fertility preservation and restoration via chemical interventions.
Current options of the tools for isolating and studying heterogeneous SSCs are still limited. THY1 can help enrich mixed A_undiff_ populations from mice [8, 35]. While studies in mice have established GFRα1 as a marker for identifying and isolating SSCs, its utility for sorting human SSCs remains scarcely explored [36, 37]. SSEA4 is absent in primitive SSCs from adult testes, thus limiting its applications [38]. The ITGA6^+^ population of human testicular cells becomes adherent following long‐term culture [39]. Here, we establish IL1R2 as a surface marker distinguishing a specific SSC subpopulation, which, despite its low native abundance, can be selectively isolated from both human and mouse samples. Critically, Il1r2 knockout males maintain normal fertility, indicating that SSC function and activity are not regulated by IL1R2 [40]. This highlights the role of IL1R2 as a non‐destructive sorting marker, preserving SSC functional integrity. In this study, we identified IL1R2^+^ SSCs in both human and mouse and explored their potential role as surface markers for cell sorting. Although SSC xenotransplantation is a major tool for assessing stem cell characteristics, we only transplanted mouse IL1R2^+^ SSCs to confirm their strong stem cell characteristics [41, 42]. Future research will explore the self‐renewal and regenerative complexities of human IL1R2^+^ SSCs through additional experiments. Although challenges remain in human SSC expansion and transplantation, IL1R2 offers a valuable tool for optimizing protocols and improving fertility restoration.
In summary, this study identifies IL1R2 as a conserved surface marker defining a distinct SSC subpopulation in both mouse and human testes. IL1R2^+^ SSCs exhibit hierarchical properties, occupy a critical developmental stage, and are essential for homeostatic spermatogenesis and regeneration. Under injury, they expand clonally and activate the PI3K‐AKT‐mTORC1 pathway, highlighting their role in A_undiff_‐mediated repair. Moreover, the ability to selectively isolate IL1R2^+^ SSCs from human and mouse testes, without impairing SSC functionality, provides a practical and non‐disruptive tool for studying SSC biology. Given the limitations of existing markers, IL1R2 represents a valuable addition for enriching functional SSCs. This work thus not only expands our understanding of SSC heterogeneity and regeneration but also offers translational promise for fertility preservation strategies, particularly in patients recovering from gonadotoxic therapies.
Conclusion
4
Spermatogonial stem cells (SSCs) provide critical support to spermatogenesis. Here, we established IL1R2 as a sorting marker for SSC, with which further insights into the SSC heterogeneity and dynamic features can be obtained. We also confirmed our findings in both mouse and human, thus validating IL1R2 as the surface marker for isolating and characterizing SSCs. Furthermore, we discovered that IL1R2^+^ SSCs can be reactive in proliferation and contribute to restoring SSCs homeostasis upon disruption, which requires the activation of the PI3K‐AKT‐mTORC1 pathway. More importantly, we confirmed that this process can be effectively enhanced by the agonist of the pathway. Overall, our findings not only provide new insights into the complexity of SSCs but also propose a promising route for isolating and preserving SSCs for therapeutic purposes.
Experimental Section
5
Ethics Approval
5.1
The adult testicular samples used for RNA sequencing analysis were obtained from males diagnosed with obstructive azoospermia (OA). All patients provided informed consent and voluntarily donated their testicular tissue for this study. The experiments conducted in this study were approved by the Ethics Committee of the Institute of Basic Medical Sciences, Chinese Academy of Medical Sciences. The study design and implementation complied with all relevant regulations regarding the use of human research participants and were conducted according to the standards set by the Declaration of Helsinki.
Animals
5.2
Il1r2‐creERT mice were generated using the endogenous Il1r2 promoter to drive creERT expression. As shown in Figure 3, creERT was inserted into the 3′ UTR region of exon 9 of the IL1R2 gene using CRISPR/Cas9. Correct insertion of the construct was verified by Southern blotting, and stable mouse lines were established by breeding the founder mice. CreERT expression was stable in IL1R2^+^ cells without affecting Il1r2 expression (Figure S4). In addition, Rosa26 ^mTmG/+^ and Rosa26 ^DTA/+^ mice were from Jackson Laboratories. For genotyping, DNA was extracted from the tail tips of the mice, and PCR was performed using specific primers (Table S1). C57/BL6 mice were used as wild‐type mice. For busulfan treatment, a single dose of 10 mg/kg busulfan was administered via intraperitoneal injection. For treatment with PI3K‐AKT‐mTORC1 agonists and inhibitors, after four days of busulfan treatment, the mice were treated with 4 mg/kg of the agonist 1,3‐Dicaffeoylquinic acid (DQA, MedChemExpress, China) [28, 29, 43, 44] and 4 mg/kg of the inhibitor rapamycin (RAPA, Selleck, China) [27] for 6 consecutive days via intraperitoneal injection. The mice were treated with 10 mg/kg of GDC‐0941 (MedChemExpress, China) [31, 32, 45] and 10 mg/kg of MHY1485 (MedChemExpress, China) [29, 30, 46] for 2 consecutive days via intraperitoneal injection. All mice were housed in a specific pathogen‐free facility under a standard 12‐h light/12‐h dark cycle with ad libitum access to water and food. The experimental procedures were approved by the Animal Care and Use Committee of the Chinese Academy of Medical Sciences and were conducted in accordance with the guidelines approved by the Animal Care and Use Committee of the Chinese Academy of Medical Sciences.
Tamoxifen Induction Method
5.3
For long‐term induction of adult Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice, they were fed a diet containing tamoxifen for four days for experiments presented in Figures 2 Jand 5H,I (Harlan Teklad, TD.130859, 400 mg/kg). For short‐term induction of adult Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice, each mouse received two consecutive dose injections of 3.0 mg each, with 12 h for experiments shown in Figures 2B,H,3A, and 5A. For low‐dose induction for clonal labeling, adult Il1r2 ^CreERT2/+^; Rosa26 ^mTmG/+^ mice received a single dose injection of 0.35 mg of 4OH‐tamoxifen dissolved in dimethyl sulfoxide (DMSO) for experiments shown in Figure 4A. After a specific pulse duration, one testis was extracted from each mouse. At the next time point, the remaining testis was extracted for analysis.
Flow Cytometry and Cell Sorting
5.4
Testicular Single‐Cell Preparation
5.4.1
After removing the tunica albuginea from the mouse testes, the testis was cleaned with PBS thrice. To remove Leydig cells and other testicular somatic cells, simple seminiferous tubules were prepared to be digested with collagenase type IV (0.5 mg/mL, Sigma) at 37°C for 5 min and cleaned thrice with PBS. The seminiferous tubules were digested at 37°C for 15 min using collagenase type IV (0.5 mg/mL, Sigma) and DNase I (0.05 mg/mL, Sigma). Digestion was terminated with serum. The resulting cell suspension was filtered through a 70‐µm cell strainer and centrifuged at 2000 rpm for 10 min. The cell pellet was resuspended in 1 mL of PBS for cell counting. For human testicular single‐cell preparation, human seminiferous tubules obtained through microdissection were first washed thrice with cold PBS and minced. The minced seminiferous tubules were digested at 37°C for 5 min using TrypLE, and the supernatant was collected. The remaining tissue pellet was digested at 37°C for 5 min using collagenase type IV and DNase I. The digestion suspensions were filtered through a 70‐µm cell strainer, and the digestion was eventually terminated with serum. The filtered cell suspension was centrifuged at 2000 rpm for 10 min at room temperature and resuspended in 1 mL of PBS for cell counting.
Flow Cytometry
5.5
The obtained single‐cell suspension was incubated with fluorochrome‐conjugated antibodies at 4°C for 30 min, with gentle mixing every 10 min. The following antibodies were used: CD90‐PE (Invitrogen, Cat: MA5‐17749, 1:200); CD117‐APC (Proteintech, Cat: APC‐65054, 1:200); E‐CADHERIN‐488 (Invitrogen, Cat: 53324982, 1:100); IL1R2‐PE (BD Biosciences, Cat: 554450, 1:50); SSEA4‐488 (for human, BioLegend, Ca: 330411, 1:50); IL1R2‐PE (for human, Invitrogen, Cat: MA5‐23543, 1:50); CD117‐APC (for human, BioLegend, Cat: 313206, 1:200); 7‐AAD (BioLegend, Cat: 420404, 1:500) or DAPI (Boster, Cat: AR1176,1:500) for dead cell exclusion. To assess non‐specific binding, the following isotype control antibodies were used in parallel at the same concentrations: Mouse IgG, Isotype Control‐PE (BioLegend, Cat: 400203); Rat IgG, Isotype Control‐PE (BD Biosciences, Cat: 553930); Rat IgG Isotype Control‐Alexa Fluor 488 (Invitrogen, Cat: 53403180); Mouse IgG Isotype Control‐Alexa Fluor 488 (BioLegend, Cat: 401323); Mouse IgG Control‐APC (BioLegend, Cat: 400119). The antibodies and their concentrations used in flow cytometry are shown in Table S1. Gating was performed based on FSC‐A and SSC‐A parameters. Doublets were removed using FSC‐H, and dead cells were excluded using 7‐AAD or DAPI. The population was gated based on fluorescence intensity, clearly distinguishable from isotype control‐stained cells. The final sorted populations were selected based on reporter gene expression or surface marker staining. Cell sorting was carried out on a BD FACSMelody Cell Sorter (BD Biosciences), and data were analyzed using FlowJo software (v10.8.1). Gating strategies are illustrated in Figure S3D. The minimum number of cells loaded per run was approximately 1 × 10⁶ total testicular cells to ensure a full representation of SSCs subpopulations, while allowing for reliable spectral unmixing and stable signal detection.
Immunofluorescence
5.6
To prepare paraffin sections, the testicular tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and cut into 7‐µm sections for paraffin dewaxing and antigen retrieval. Sections were blocked with 3% BSA at room temperature for 1 h. After incubating with the primary antibody overnight at 4°C, the sections were washed thrice with PBS. Next, the sections were incubated with the secondary antibody at room temperature for 1 h. After three washes with PBS, the coverslips were mounted on glass slides with an anti‐fade mounting medium containing DAPI for fluorescence quenching.
For whole‐mount immunostaining of seminiferous tubules, tubules were fixed in 4% paraformaldehyde for 1 h, followed by three washes with PBS. Tubules were blocked with 3% BSA for 1 h, incubated with primary antibody overnight at 4°C, and eventually washed with PBS. Next, the tubules were incubated with the secondary antibody for 2 h. Finally, the tubules were mounted on glass slides using an anti‐fade mounting medium containing DAPI for fluorescence quenching. The images were captured using a confocal microscope (Leica).
The following antibodies were used for immunostaining: anti‐IL1R2 (with dilution by 1:100, Invitrogen, PA5‐47759), anti‐GFRα1 (0.5 mg/ml, R&D, AF560), anti‐SOX3 (1:300, GeneTex, GTX129235), anti‐ZBTB16 (1:200, Santa Cruz, sc‐28319), anti‐C‐KIT (1:200, R&D, AF1356, MAB1356), anti‐LIN28A (1:300, Abcam, ab63740), anti‐EOMES (1:300, Abcam, ab23345), anti‐PAX7 (1:300, Developmental Studies Hybridoma Bank), anti‐MKI67 (1:300, Abcam, ab279653), anti‐FOXC2 (1:300, R&D, AF6989). Secondary antibodies were Alexa Fluor‐conjugated from Thermo or Jackson ImmunoResearch and used at 1:500 dilutions. The antibodies and their concentrations are listed in Table S1.
Histological Examination
5.7
Prior to staining, paraffin‐embedded testicular tissue sections were subjected to three rounds of dewaxing in xylene, followed by rehydration with different concentrations of ethanol. Subsequently, the sections were stained with hematoxylin and eosin. After staining, the sections were dehydrated in ethanol and xylene and fixed with neutral resin. Finally, the sections were observed under a microscope.
Total RNA Isolation and Quantitative Real‐Time PCR
5.8
The isolated cells were lysed in TRIzol reagent to extract total RNA from whole cells, and the RNA was quantified using a NanoDrop 2000 spectrophotometer. Complementary DNA (cDNA) was synthesized using a cDNA synthesis kit. qPCR experiments were conducted using SYBR Green, and the data were normalized to the expression levels of a housekeeping gene (Gapdh). Relative gene expression was quantified using the double‐delta Ct method. The primers used for qPCR are listed in Table S1.
RNA Sequencing
5.9
Cells sorted and purified by flow cytometry were subjected to total RNA extraction, followed by quantification using a NanoDrop 2000. The Illumina RNA‐Seq protocol was used to construct standard RNA‐Seq libraries. The libraries were then sequenced on the Illumina NovaSeq platform, generating paired‐end reads with a length of 150 bp. The raw FASTQ format data were filtered and trimmed using Trimmomatic and mapped to the complete mouse reference genome using STAR. The read counts aligned to genes were calculated using featureCounts, followed by the analysis of differentially expressed genes using the DESeq2R package. Enrichment analysis of Gene Ontology (GO), Kyoto Encyclopedia of Genes and Genomes (KEGG), and Gene Set Enrichment Analysis (GSEA) was performed using the clusterProfiler package.
Single‐Cell Sequencing Analysis
5.10
Raw sequencing data were processed using the Cell Ranger pipeline with default parameters. The resulting gene‐barcode matrices were further analyzed using the Seurat package (v4.0) in R. Low‐quality cells were filtered out based on the following criteria: cells with <200 or >6,000 detected genes, or <10% mitochondrial gene content were excluded. Genes expressed in fewer than three cells were also removed. After normalization using LogNormalize, the top 2,000 highly variable genes were identified using the vst method for downstream analysis. Principal component analysis (PCA) was performed on the variable genes, and the top 20 PCs were used for clustering using the FindNeighbors and FindClusters functions. Uniform Manifold Approximation and Projection (UMAP) was used for dimensionality reduction and visualization. Cluster identities were assigned based on canonical marker gene expression and differential gene analysis using FindAllMarkers (logfc. threshold = 0.25, adjusted p < 0.05). FeaturePlot and VlnPlot functions were used to visualize the specific genes. For pseudotime trajectory analysis, cells from undifferentiated spermatogonia clusters were subset and analyzed using Monocle. The trajectory graph was learned using default parameters and ordered along pseudotime based on transcriptional progression inferred from cluster 0 (primitive SSC state). Differential gene expression along the trajectory was calculated using graph_test with Moran's I statistic, and temporally dynamic genes were clustered into modules for interpretation. Highly variable genes specific to early SSC‐enriched clusters (cluster 0) were selected as candidate gene sets for downstream surface marker screening. Gene Ontology (GO) and Gene Set Enrichment Analysis (GSEA) analysis was performed using the clusterProfiler package (v4.2) (adjusted p < 0.05).
SSC Culture
5.11
The procedure of obtaining single cells from 7‐day‐old mouse testes and flow cytometry‐based sorting of IL1R2^+^ cells was conducted as described previously [47]. The sorted cells were cultured on mouse embryonic fibroblasts treated with mitomycin C. The cells were cultured in a mouse SSC serum‐free medium supplemented with recombinant human GDNF (20 ng/mL), recombinant human FGF (1 ng/mL), and other components. The culture was maintained at 37°C with 5% CO_2_ in a humidified incubator. The medium was changed every two days, and passaging was performed every 7 days.
Microinjection Transplantation
5.12
Prior to the transplantation experiment, 8‐week‐old C57 wild‐type mice, which served as recipients, were treated with a high dose of busulfan (40 mg/kg). After one month, an equal number of cells (1 × 10^4^ cells per testis) were injected into the seminiferous tubules of recipient mice using a microinjection technique. Two months after transplantation, the recipient testes were observed and photographed under a fluorescence microscope. Fluorescent colonies were defined as those occupying more than 50% of the basal surface of the seminiferous tubules and having a length of at least 0.1 mm. The number of colonies in the recipient testes represents the quantity of functional SSCs.
Statistical Analysis
5.13
As shown in the legend, the significance of differences in the data was analyzed using a non‐paired two‐tailed Student's t‐test. All results are presented as mean values with standard error of the mean (SEM), and the sample size is indicated in each legend. Significance is denoted as non‐significant (ns), ^^ p < 0.05, ^^ p < 0.005, ^^ p < 0.001, and ^****^ p < 0.0001. No statistical method was used to determine the sample size.
Author Contributions
P.L. and W.S. conceived and designed the study; P.L. performed most of the experiments and analyzed the data with the help of Z.Z., Z.X., and C.J.; Z.W., J.T., M.L., D.Z., X.M., D.C., Q.G., C.Y., J.Z., Y.L.and J.P. provided additional experimental support; S.M., N.Z., J.P., K.L., and W.S. provided critical suggestions on manuscript preparation; P.L., K.L., and W.S. wrote the manuscript with help from all authors.
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Supporting File: advs73423‐sup‐0001‐SuppMat.docx.
Supporting File: advs73423‐sup‐0002‐Supplementary Table1.xlsx.
Supporting File: advs73423‐sup‐0003‐Supplementary Table2.xlsx.
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