All animal studies were approved by the Institutional Animal Care and Use Committee of Washington University School of Medicine, Saint Louis, MO, USA and Use Committee of Baylor College of Medicine, Houston, TX, USA. Atg14 f/f mice were provided by gift from Dr. Shizuo Akira at the Department of Host Defense, Research Institute for Microbial Diseases (RIMD), Osaka University, and previously described [28]. Wild type Pr-cre mice were provided by Dr. John Lydon at Baylor College of Medicine, Houston and previously described [29]. Atg14^flox/flox mice, in which exon 4 was flanked by loxp sites, were bred to progesterone receptor cre (PR^cre/+) mice to generate (Atg14^flox/flox; PR^cre/+ mice), hereafter referred to as Atg14 cKO mice. Both control and conditional knockout females were generated by crossing females carrying homozygous Atg14^flox/flox alleles with Atg14 cKO males. Foxj1-cre mice were a generous gift from Dr. Michael J. Holtzman at Washington University St. Louis and previously described [30]. Foxj1/Atg14 cKO mice were generated by crossing females carrying homozygous Atg14 ^f/f/Foxj1^f/f females with Atg14^f/f/Foxj1-cre males. All mice were age-matched and on a C57BL/6 genetic background (The Jackson Laboratory, Bar Harbor, ME). Mice were genotyped by PCR analysis of genomic DNA isolated from tail clippings using the gene-specific primers listed in Supplementary Table 1.

Female fertility was determined by mating cohorts of Atg14 cKO experimental (n=6) and control Atg14 f/f (n=4) females individually starting at 8 weeks of age with sexually mature males of proven fertility. Similarly, breeding trials were set up for Foxj1/Atg14-cre females (n=6) and Foxj1/Atg14 control 8-week-old f/f females (n=6). The numbers of litters and pups were tracked over 6 months for each female. Pups per litter for each genotype are reported as mean ± SEM. For timed mating, the morning on which the copulatory plug was first observed was considered 1 dpc. To visualize implantation sites, mice received a tail vein injection of 50 μL of 1% Chicago Sky Blue dye (Sigma-Aldrich, St. Louis, MO, USA) at 5 dpc just before sacrifice.

The hormonal profile of pregnancy at the time of implantation was done using a previously described experimental scheme [26]. Briefly, Atg14 cKO and control females (8 weeks old) were bilaterally ovariectomized under ketamine anesthesia with buprenorphine-SR as an analgesic. Mice were allowed to rest for two weeks to dissipate all endogenous ovarian hormones. After the resting period, mice were injected with 100 ng of estrogen (E2; Sigma-Aldrich) dissolved in 100 µL of sesame oil on two consecutive days and then allowed to rest for two days. At this point, mice were randomly divided into three groups of five: Vehicle-treated (E2 priming) mice received four consecutive days of sesame oil injections; E2 group mice received three days of sesame oil injections followed by a single injection of 50 ng of E2 on the fourth day; The E2/P4 mice received 1 mg of progesterone (P4; Sigma-Aldrich) for three consecutive days followed by a single injection of 1 mg P4 plus 50 ng E2 on the fourth day. All hormones were delivered by subcutaneous injection in a 90:10 ratio of sesame oil: ethanol. Mice were euthanized 16 hours after the final hormone injection to collect the uteri. A small piece of tissue from one uterine horn was processed in 4% neutral buffered paraformaldehyde for histology, and the remaining tissue was snap-frozen and stored at −80°C.

For serum hormone levels measurement, blood was collected from D4 pregnant mice before mice were sacrificed. Serum was separated from the blood by centrifugation and stored at −80°C before hormone analysis. Serum P4 and E2 levels were measured by using ELISA kits (Enzo life Sciences) according to the manufacturer’s instructions.

Tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and then sectioned (5 μm) with a microtome (Leica Biosystem, Wetzlar, Germany). Tissue sections were deparaffinized, rehydrated, and stained with Hematoxylin and Eosin (H&E) as described previously [31]. All the histology was performed on three sections from each tissue of individual mice, and one representative section image is shown in the respective figures.

For histological analysis, the collected tissues (oviduct or uteri) were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned. Sections (5 µm) were immunostained (n=5 per group) as described previously [31]. Briefly, after deparaffinization, sections were rehydrated in an ethanol gradient and then boiled for 20 min. in citrate buffer (Vector Laboratories Inc., Newark, CA, USA) for antigen retrieval. Endogenous peroxidase activity was quenched with Bloxall (Vector Laboratories Inc.), and tissues were blocked with 2.5% goat serum in PBS for 1 hr (Vector Laboratories Inc.). After washing in PBS three times, tissue sections were incubated overnight at 4 °C in 2.5% goat serum containing the primary antibodies listed in Supplementary Table 2. Sections were incubated for 1 hr with biotinylated secondary antibody, washed, and incubated for 45 min with ABC reagent (Vector Laboratories Inc.). Color was developed with 3, 3’-diaminobenzidine (DAB) peroxidase substrate (Vector Laboratories Inc.), and sections were counter-stained with hematoxylin. Finally, sections were dehydrated and mounted in Permount histological mounting medium (Thermo Fisher Scientific, Waltham, MA, USA).

For ultrastructural analysis, oviducts were fixed in 2% paraformaldehyde/2.5% glutaraldehyde (Ted Pella Inc., Redding, CA) in 100 mM cacodylate buffer, pH 7.2 for 1 hr at room temperature and then overnight at 4°C. Samples were washed in cacodylate buffer and postfixed in 1% osmium tetroxide (Ted Pella Inc.) for 1 hr. Samples were then rinsed extensively in dH20 prior to en bloc staining with 1% aqueous uranyl acetate (Ted Pella Inc.) for 1 hr. Following several rinses in dH20, samples were dehydrated in a graded series of ethanol and embedded in Eponate 12 resin (Ted Pella Inc.). For initial evaluation semithin sections (0.5 μm) were cut with a Leica Ultracut UCT7 ultramicrotome (Leica Microsystems Inc., Bannockburn, IL) and stained with toluidine blue. Sections of 95 nm were then cut and stained with uranyl acetate and lead citrate and viewed on a JEOL 1200 EX II transmission electron microscope (JEOL USA Inc., Peabody, MA). Images at magnifications of 3,000X to 30,000X were taken with an AMT 8-megapixel digital camera (Advanced Microscopy Techniques, Woburn, MA).

Formalin-fixed and paraffin-embedded sections were deparaffinized in xylene, rehydrated in an ethanol gradient, and boiled in a citrate buffer (Vector Laboratories Inc.) for antigen retrieval. After blocking with 2.5% goat serum in PBS (Vector laboratories) for 1 hr at room temperature, sections were incubated overnight at 4°C with primary antibodies (Supplementary Table 2) diluted in 2.5% normal goat serum. After washing with PBS, sections were incubated with Alexa Fluor 488-conjugated secondary antibodies (Life Technologies, Carlsbad, CA, USA) for 1 hr at room temperature, washed, and mounted with ProLong Gold Antifade Mountant with DAPI (Thermo Fisher Scientific). All Immunofluorescence images were obtained using a Zeiss LSM 880 confocal microscope (10x and 40x objective lens).

Fresh oviduct tissues from 8-week-old mice from control (n=3) and cKO (n=3) were collected and fixed in 4 % PFA at 4°C overnight. After that, oviducts were washed with PBS and cryoprotected using gradually increased sucrose concentrations in a row (15 and 30% w/v). 24 h later, oviducts were embedded in an OCT medium, frozen (−80 °C), and cryo-sectioned in the CM3050S Cryostat (Leica Biosystems, Germany). The cryosections were mounted on Superfrost plus glass slides (Fisher Scientific, Pittsburgh, PA) and stored at −80°C until used.

Cryo-sections were processed for immunofluorescence staining as described before [32]. Briefly, cryosections were blocked in 2.5% normal goat serum. After washing with PBS, sections were blocked with Mouse on Mouse (M.O.M.) IgG blocking reagent (M.O.M. Fluorescein Kit, Vector Laboratories, #FMK-2201) diluted in 1% BSA per the manufacturer’s instructions. After blocking, sections were incubated overnight at 4°C with primary antibodies (Supplementary Table 2) diluted in 2.5% normal goat serum. Next day, following washing with PBS, sections were incubated with Alexa Fluor 488-conjugated secondary antibodies (Life Technologies, Carlsbad, CA, USA) for 1 hr at room temperature, washed, and mounted with ProLong Gold Antifade Mountant with DAPI (Thermo Fisher Scientific). All Immunofluorescence images were imaged using a Nikon Fluorescent microscope.

Protein lysates (40 µg per lane) from uteri or oviducts were loaded on a 4-15% SDS-PAGE gel (Bio-Rad, Hercules, CA, USA), separated in 1X Tris-Glycine Buffer (Bio-Rad), and transferred to PVDF membranes via a wet electro-blotting system (Bio-Rad), all according to the manufacturer’s directions [33]. PVDF membranes were blocked for 1 hour in 5% non-fat milk in Tris-buffered saline containing 0.1% Tween-20 (TBS-T, Bio-Rad), then incubated overnight at 4 °C with antibodies listed in Supplementary Table 2 in 5% BSA in TBS-T. Blots were then probed with anti-Rabbit IgG conjugated with horseradish peroxidase (1:5000, Cell Signaling Technology, Danvers, MA, USA) in 5% BSA in TBS-T for 1 hr at room temperature. Signal was detected with the Pierce™ ECL Western Blotting Substrate (Millipore, Billerica, MA, USA), and blot images were collected with a Bio-Rad ChemiDoc imaging system.

Tissues/cells were lysed in RNA lysis buffer, and total RNA was extracted with the Purelink RNA mini kit (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. RNA was quantified with a Nano-Drop 2000 (Thermo Fisher Scientific). Then, 1 µg of RNA was reverse transcribed with the High-Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). The amplified cDNA was diluted to 10 ng/µL, and qRT-PCR was performed with primers listed in the Supplementary Table 1 and TaqMan 2X master mix (Applied Biosystems/Life Technologies, Grand Island, NY, USA) on a 7500 Fast Real-time PCR system (Applied Biosystems/Life Technologies). The delta-delta cycle threshold method was used to normalize expression to the reference gene 18S.

Polyphyllin VI (Selleck chemicals, Houston, TX, USA), a pharmacological agent that induces caspase-1-mediated pyroptosis, was used to study its effects on embryo transport in the oviduct [34]. Eight-week-old C57BL/6 mice were injected for three consecutive days starting from 1 dpc with Polyphyllin VI activator dissolved in 40% PEG-300 (15mg/kg body weight). Dimethyl sulfoxide with 40% PEG-300 was administered as a vehicle.

A two-tailed paired student t-test was used to analyze data from experiments with two experimental groups and one-way ANOVA followed by Tukey’s post hoc multiple range test was used for multiple comparisons. All data are presented as mean ±SEM. GraphPad Prism 9 software was used for all statistical analyses. Statistical tests, including p values, are reported in the corresponding figure legends or, when possible, directly on the data image. To ensure the reproducibility of our findings, experiments were replicated in a minimum of three independent samples, to demonstrate biological significance, and at least three independent times to ensure technical and experimental rigor and reproducibility.

To explore the role of Atg14 in uterine function, we first determined its expression levels in uterus during early pregnancy (Day 1-7) in mice. We found distinct expression of ATG14 in all the uterine compartments (luminal epithelium, glands, and stroma) by day 1 of pregnancy, which disappeared by day 2 of pregnancy (Fig. 1A). However, the ATG14 expression in uterus reappeared by day 3 and persisted through the day 7 of pregnancy. The period from day 3 to day 7 is critical, as it is during this time when uterus begins to prepare for embryo implantation and undergoes decidualization process. Consistent with protein expression, similar Atg14 expression at mRNA levels was noted in uteri from early pregnant mice (Fig. 1B). This analysis suggests a potential role for ATG14 protein in the uterine physiologic adaptations during early pregnancy. Thus, to study the role of ATG14 in uterine function, we generated a conditional knockout (PR^cre/+ /Atg14^flox/flox) mouse model by crossing Atg14 flox/flox mice with mice expressing Cre recombinase under the control of progesterone receptor promoter (PR^cre/+). Histological examination of the uterus from adult females showed no gross morphological differences between Atg14 cKO and control mice (Fig. S1A). Further, we did not find any overt defects in ovary as cKO mice had normal follicles and corpus luteum as like their corresponding controls (Fig. S1B). Analysis of transcript levels from the uterus, ovary, and liver showed that while Atg14 levels were efficiently depleted in uteri from cKO mice the levels were unaltered in the ovary and liver samples in control and cKO mice (Fig. 1C). Immunofluorescence analysis further confirmed the efficient deletion of ATG14 in all uterine compartments (epithelium and stroma) of cKO mice uteri compared to corresponding control groups (Fig. 1D).

Considering the effective ablation, a 6-month breeding study was performed mating virile wild-type male mice with adult Atg14 cKO and control female mice. We found that Atg14 cKO females did not deliver any litter during the 6-months trial period. However, the control female (Atg14 flox/flox) mice delivered an average of ∼7-8 pups per litter every month (Table 1) (Fig. 1E). These findings suggest an indispensable role of Atg14 in female fertility with intact ovarian function.

Based on the normal ovarian morphology, we posited that the infertility observed in females with Atg14 cKO status could be attributed to compromised uterine functioning. Analysis of the implanting embryos at Day 5 of pregnancy showed no implantation sites in the uteri of Atg14 cKO females, whereas ∼8 to 9 implantation sites were seen in the uteri of control mice at 5 dpc (Fig. 2A, left panel). Whilst there was no embryo present in the uterine lumen of Atg14 cKO mice, a fully attached embryo encapsulated by the luminal uterine epithelium, was seen in control mice uteri (Fig. 2A, middle panel). Additionally, MUC1, a receptive marker expression persisted in the luminal epithelium of cKO mice uteri at 5 dpc as well as 4 dpc (Fig. 2A, right panel, Fig. 2C). The process of successful embryo attachment and implantation within the uterus necessitates a transition from a non-receptive to a receptive state, a transformation orchestrated under the regulated influence of steroid hormones. Thus, the effects of Atg14 loss on uterine responsiveness to steroid hormones E2 and P4 were characterized. We performed an established hormone-induced uterine receptivity experiment, which involved ordered and co-stimulatory actions of E2 and P4 leading to the initiation of a receptive phase [35]. In response to E2 treatment, uteri from both Atg14 control and cKO mice showed similar proliferation as evidenced by Ki-67 staining (Fig. 2B, middle panel). Following the co-stimulation with E2+P4 treatment, epithelial proliferation was inhibited with concomitant induction of stromal cell proliferation in control mice uteri indicating fully receptive uteri (Fig. 2B). Interestingly, cKO mice uteri failed to elicit sub-epithelial stromal cell proliferation and showed intact P4-driven inhibition of epithelial proliferation. Consistently, uteri from Atg14 cKO mice at 4 dpc showed a reduced number of proliferating stromal cells (Fig. 2D). The absence of significant changes in E2-induced targets, such as Lif, Mcm2, Ccnd1, and Fgf18, at 4 dpc (Fig. 2E) supports our conclusion that ATG14 is required for P4-mediated but not for E2-mediated actions during uterine receptivity. Moreover, the normal serum levels of E2 and P4 at D4 of pregnancy rule out any hormonal imbalances, strongly suggesting that the observed phenotype is primarily due to the uterine-specific loss of Atg14 (Fig. 2F).

Given that Atg14 cKO mice had impaired embryo implantation, we wondered whether embryos are reaching the uterus timely through the oviduct. To determine this, we flushed embryos from both the control and cKO mice uteri on day 4 of pregnancy. Interestingly, in cKO mice, we could retrieve only 1-2% of embryos from their uteri, whereas in control mice, 100% of well-developed blastocysts were retrieved from their uteri (Fig. 3A & B). To ensure the timely transport of all embryos from the oviducts to the uteri, we also flushed oviducts from both control and cKO mice. As expected, in the control mice oviduct flushing, we could not recover any embryos or blastocysts, indicating their precise and timely transport to the uterus. Unexpectedly, oviduct flushing from cKO mice resulted in the retrieval of approximately 80-90% of blastocysts, suggesting their potential entrapment within the oviducts, impeding their transit to the uterus (Fig. 3A & B). At 4 dpc, there was no significant difference in the average number of blastocysts, morula, or non-viable or retrieved from Atg14 control uteri and cKO oviducts (Fig. S4A). However, we noted that the percentage of developmentally delayed embryos appeared to be higher in Atg14 cKO oviducts compared to the embryos retrieved from control uteri (Fig. 3C). The histological analysis further confirmed an entrapped embryo in the ampulla of Atg14 cKO at 4 dpc as shown in Fig. 3D. Given the embryo retention phenotype in oviduct, we sought to determine if ATG14 is expressed in this region. Consistent with previous studies reporting PR-cre activity in the isthmus [8, 29], we found a significant depletion of ATG14 in the isthmus compared to the ampulla (Fig. S2A). Further, the increased p62 and LC3B expression in cKO oviducts suggests that the observed embryo retention phenotype might be attributed to the loss of ATG14-dependent autophagy in cKO oviducts (Fig. S2B).

Similarly, when we super-ovulated the mice and looked for the embryos on day 4 of pregnancy, we could recover ∼20-25 embryos from the cKO mice oviducts compared to only 4-5 embryos that were able to reach the uterus. In comparison, in control mice, 90% of embryos were able to complete their pre-destined and timely transport to the uterus (Fig. 3E), except 1-2% of unfertilized embryos, which remained in their oviducts.

To understand the underlying cause for retained embryos in cKO mice oviducts, we performed histology analysis and determined the structural morphology of their oviducts. The cKO mice oviduct lining shows marked eosinophilic cytoplasmic change akin to decidualization in human oviducts. Some of the cells showed degenerative changes with cytoplasmic vacuolization and nuclear pyknosis, loss of nuclear polarity, and loss of distinct cell borders giving an appearance of fusion of cells (Fig. 3F). The marked cytologic enlargement appears to cause luminal obliteration or narrowing resulting in a completely unorganized, obstructed, and narrow lumen, thereby, hampering the path of embryos to reach the uterus as evident from an entrapped embryo shown in Fig. 3D. Further, epithelial (cytokeratin KRT8-positive) and myosalpinx (α-smooth muscle active [SMA]-positive,) marker analysis revealed completely distorted epithelial structures (in terms of loss of epithelial cell integrity) with no overt defects in the muscle organization in oviducts of cKO mice (Fig. 3F, lower panel). Notably, the ampullary region from cKO oviducts seems to be normal with intact epithelial and smooth muscle structures as evident from KRT8 and SMA staining (Fig. 3F, upper panel).

The oviduct epithelium primarily consists of two types of cells: ciliated and secretory (non-ciliated) cells. Ciliated cells play a role in embryo and oocyte transport by means of ciliary beat, and non-ciliated/secretory cells produce an oviductal fluid that is rich in amino acids and various molecules, thereby providing an optimal micro-environment for sperm capacitation, fertilization, embryonic survival, and development [36]. Therefore, we determined whether oviducts from cKO mice possess normal ciliated and secretory cell composition. To do so, we examined the expression of various markers, including acetylated-α-tubulin (cilia marker), FOXJ1 (ciliogenesis markers), and PAX8 (non-ciliated/ secretory cell marker) in control and cKO oviducts. As shown in Fig. 4A, while we observed normal ciliary structures in the ampulla of both control and cKO oviducts, there was a substantial loss of the ciliary epithelial cells (indicated by fewer α-tubulin and FOXJ1-positive cells) (Fig. 4B, left panel and Fig. S3) as well as secretory cells (indicated by fewer PAX-8 positive cells) in the isthmus of cKO oviducts (Fig. 4B, right panel).

Given the importance of cilia in embryo transport, we wondered whether the loss of Atg14 in oviduct cilia has any impact on embryo transport. To address this, we generated a Foxj1-cre/+; Atg14 f/f mouse model, wherein Atg14 will be ablated only in ciliary epithelial cells. We observed that ciliated epithelial cells that were positive for acetylated α-tubulin staining did not appear to be different in Foxj1^Cre/+; Atg14 f/f mice oviduct compared to controls suggesting normal ciliogenesis in Foxj1^Cre/+; Atg14 f/f mice (Fig. 4C).

The 6-month breeding trial revealed that loss of Atg14 in oviductal cilia had no impact on fertility (Table 2, Fig. 4D & E). Consistently, D5 implantation study analysis showed ∼7-8 visible embryo implantation sites in Foxj1/Atg14 cKO mice like their corresponding controls suggesting that embryos were able to make their way to the uterus in a timely manner and undergo implantation despite the ablation of Atg14 in the oviduct ciliary epithelial cells (Fig. 4F). These findings suggest that ciliary expression of Atg14 is dispensable for embryo transport.

To gain more insights into structural defects, we performed TEM analysis on oviducts collected from both control and cKO females on day 4 of pregnancy. Interestingly, oviducts from cKO females had numerous altered mitochondrial structures with abnormally enlarged mitochondria and a less dense matrix compared to control oviducts that had small and compact mitochondria with tight cristae and a dense matrix (Fig. 5A). Corroborating and extending those findings, we observed marked reductions of mitochondria network (as defined by TOMM20 staining) in the sub-nuclear region of oviducts from Atg14 cKO mice compared to more densely packed mitochondrial network in the sub-nuclear region of control ones (Fig. 5B). Further co-localization study analysis revealed a uniform co-localization of Cytochrome C with the TOM20 positive mitochondrial network in control oviducts reflecting the intact mitochondrial integrity (Fig. 5B). In contrast, in Atg14 cKO oviducts, cells with disrupted mitochondrial networks exhibited increased cytosolic leakage of Cytochrome C (Fig. 5B). Additionally, analysis of various mitochondrial functional and architecture markers (Cox4i2, Pink1, Opa1, and Drp1 showed reduced expression in Atg14 cKO compared to controls (Fig. 5C).

Having identified the cellular mechanisms of Atg14, we aimed to delineate the underlying molecular mechanisms associated with Atg14 function in the oviduct. Since we found mitochondrial defects with ATG14 loss and altered mitochondrial structures have been linked to the activation of pyroptosis, we determined if ATG14 regulates pyroptosis in oviduct [37–39]. First, we assessed the key primary executors of the pyroptosis pathway, Gasdermin D (GSDMD), and caspase-1. Immunofluorescence analysis revealed a remarkable upregulation in GSDMD and caspase-1 expression in the cKO oviducts compared to controls (Fig. 5D & E). However, histological analysis of uterus and ovary showed no induction in GSDMD expression compared to their corresponding controls (Fig. 5F). Western blot analysis further confirmed elevated expression of caspase-1 and GSDMD in cKO oviducts in comparison to control oviducts as shown in Fig. 5G. Additionally, the qPCR analysis demonstrated elevated levels of inflammatory markers, such as Tnf-α and Cxcr3 in cKO oviducts compared to control ones (Fig. 5H). Based on these findings, we posit that Atg14 plays a crucial role in regulating the pyroptotic pathway by preserving the mitochondrial structural and functional integrity, and activation of pyroptosis owing to loss of ATG14 within the oviduct.

To substantiate the notion, we also evaluated the impact of unscheduled activation of pyroptosis on embryo transport. To test this, we employed a pyroptosis inducer, Polyphyllin VI, and chose the optimal dose based on the established studies [34]. Wild-type females were mated with virile males and following the plug detection treated with Polyphyllin VI for three consecutive days from 1 to 4 dpc (Fig. 5E). We chose to treat to 1 to 4 dpc for treatment to activate unwarranted pyroptosis in oviduct during the embryo transport. Following the treatments, the oviducts, and uteri from both vehicle-treated and Polyphyllin VI-treated were flushed. We found that pregnant females treated with Polyphyllin VI showed ∼50% embryo retention in the oviduct, whereas in the vehicle-treated group, no embryos were retained in the oviduct (Fig. 6A-C). Further analysis of embryos retrieved from Polyphyllin-treated oviducts showed more percentage of developmentally delayed and non-viable embryos compared to embryos recovered from control uteri (Fig. 6D). The average number of embryos recovered from polyphyllin- and vehicle-treated mice was not significantly different (Fig. S4B). Histological analysis showed a marked induction in GSDMD expression compared to vehicle-treated mice oviducts (Fig. 6E). However, precisely activating the unscheduled pyroptosis during the critical period of embryo transport is technically challenging. Nonetheless, our findings provide evidence that unscheduled pyroptosis adversely affects embryo transport through the oviduct. Taken together, these results demonstrate that Atg14 safeguards pyroptosis activation in the oviduct and allows the smooth transport of embryos to the uterus (Fig. 6F).