While ABHD2 has been described as an important regulator of sperm function (Miller et al., 2016), the function of ABHD2 in female reproduction was not known. Interestingly, unlike the male reproductive tissues, we found that ABHD2 is not detected in the female gametes but is predominantly expressed in the stromal cells surrounding the developing follicles (Figure 1a). The presence of both ABHD2 protein and mRNA was observed in pre-pubertal mouse ovaries, as shown here by immunohistochemical (IHC) staining and reverse transcriptase quantitative PCR (RT-qPCR; Figures 1b,c). Around 1 month after birth, when the ovulatory cycle begins, ABHD2 expression is further found in the lutein cells of CL (Figure 1a). In correlation with this, qPCR studies showed highest Abhd2 mRNA presence right after ovulation during the estrus stage (Figure 1c), and the expression levels are even further increased after induction of superovulation by gonadotropin injection (Figure 1c).

To study the role of ABHD2 in female reproduction, we have generated an Abhd2 KO mouse line by utilizing the CRISPR-EZ technique (Chen et al., 2016), where single guide RNA (sgRNA)/Cas9 complexes are delivered into mouse zygotes by electroporation. Cas9-mediated deletion of Abhd2 exon 6 was performed through intron-specific binding of two sgRNAs (Figure 2a). The deletion resulted in the removal of a serine (S208), the amino acid important for the catalytic function of ABHD2, and resulted in a frameshift and a premature stop codon in exon 7 (Supplementary Figure 2). Complete ablation of Abhd2 was further confirmed by genotyping PCR (Figure 2b), qPCR (Figure 2c), Western blotting (Figure 2d), and IHC analysis of the mouse ovary (Figures 2e,f). The KO mice were born at a Mendelian ratio when breeding heterozygous males with females. No obvious morphological or health phenotypes have been observed, and both homozygous and heterozygous females appeared fertile and healthy with similar body weight as their wild-type littermates (Supplementary Figure 3A).

Both the litter size and birth rate of Abhd2^+/– and Abhd2^–/– females were not significantly different from that of wild-type mice (Table 1). In addition, spontaneous ovulation gave rise to similar numbers of eggs for all genotypes (Figure 3A). However, when injected with gonadotropins, pregnant mare serum gonadotropin (PMSG) and human chorionic gonadotropin (hCG)—a standard method to induce superovulation (Christenson and Eleftheriou, 1972), both the Abhd2^+/– and Abhd2^–/– females produced significantly higher numbers of ovulated eggs (Figure 3B). The phenotype was not a result of prematurely ovulated follicles, as in vitro fertilization (IVF) studies led to a similar success rate in formation of blastulae as when using eggs from wild-type females (Abhd2^+/+: 72.4% ± 11.3% n = 3, Abhd2^+/–: 81.5% ± 9.1%, Abhd2^–/–: 75.0% ± 8.3% fertilized eggs, n = 4 for both genotypes). The fertility of 7- to 8-month-old Abhd2^+/– and Abhd2^–/– females in normal breeding was similar to that of wild-type mice (Table 1). Superovulation of the older animals up to 15-month old resulted in a similar decline in the number of ovulated eggs, although some heterozygous and homozygous mice still showed numbers comparable with that of the much younger animals (Figure 3C).

Interestingly, when following the estrous cycle of virgin females for 1 month, the Abhd2^–/– females presented with a significantly higher percentage of days in estrus, while proestrus and diestrus were shortened compared with the wild-type females (Figures 3D-F and Supplementary Figure 4). Although the Abhd2^–/– mice showed a prolonged estrus, they cycled in a regular fashion (Abhd2^+/+: 4.75 ± 0.33 days/cycle, Abhd2^+/–: 4.8 ± 0.53 days/cycle, Abhd2^–/–: 4.43 ± 0.29 days/cycle) and gave rise to litters at a similar rate as Abhd2^+/+ females (Table 1). Previous studies have shown that altered estrogen and androgen levels during pregnancy can affect the estrous cycle of female pups (Abbott et al., 2006). To determine whether or not the maternal genotype influenced the observed phenotype, the estrous cycle of Abhd2^+/– females born from wild-type mothers who were mated with homozygous males or homozygous mothers mated with heterozygous males were compared. The month-long measurements showed a similar prolonged estrus in all females (Abhd2^+/– females born from Abhd2^+/+ mothers: 47.2% ± 3.7%; Abhd2^+/– mothers: 46.5% ± 6.1%; Abhd2^–/– mothers: 46.7% ± 1.9% of days in estrus), while wild-types had only 35.6% ± 5.6% of days in estrus. These results indicate that the observed phenotype was mainly due to the genotype of the pups and not due to epigenetic influence.

To determine whether or not the increased number of ovulated eggs produced during superovulation was due to an altered follicular development, early antral and antral follicles and the corpora lutea of ovaries from Abhd2^+/+, Abhd2^+/–, and Abhd2^–/– littermate females in proestrus were counted. There was a significant decrease in the number of early antral follicles of both Abhd2^+/– and Abhd2^–/– ovaries compared with those of Abhd2^+/+ mice (Figure 4a). Instead, the females showed an increased number of antral, pre-ovulatory, follicles, and, in the case of Abhd2^+/– females, a higher number of corpora lutea (Figure 4a). However, these differences did not lead to an altered weight of the ovaries when comparing the different genotypes (Supplementary Figure 3B). Hematoxylin staining of the ovary showed a difference in atretic versus healthy, living follicles, with a larger number of atretic follicles present in Abhd2^+/– and Abhd2^–/– ovaries compared with those of Abhd2^+/+ mice (Figures 4b,c). This was further confirmed by terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) staining, where follicles going through atresia showed increased labeling of granulosa cells (Figures 4d,e). When counting only the healthy antral follicles, no genotype-specific difference in follicle number was observed. Although the Abhd2^+/– mice displayed increased follicle atresia compared with the wild-type mice, the number of healthy antral follicles was comparable with that of Abhd2^+/+ ovaries (Figure 4b).

The increased transition from early antral to antral follicles after Abhd2 ablation could account for the observed phenotype of superovulated Abhd2^+/– and Abhd2^–/– females. However, as progesterone is needed to initiate the signaling pathways leading to ovulation, in vitro follicle culture was performed to determine the role of ABHD2 in this process. Follicles from immature mice were collected and cultured for 4 days, after which ovulation was induced by the addition of hCG to the culture media (Figures 5A–C). The percentage of ovulated follicles was similar between Abhd2^+/+ and Abhd2^–/– females (Figure 5A). However, as the initial number of large follicles (240 ± 10 μm in diameter) collected from the Abhd2^–/– ovaries was higher than that from the wild-type ovaries, the total number of ovulated follicles was increased in cultures from the Abhd2^–/– females (Figure 5B). These results are similar to the higher number of ovulated eggs observed after superovulation of Abhd2^–/– mice. In conclusion, Abhd2 ablation does not appear to affect the ovulation process directly but rather gives rise to an increased number of mature follicles, possibly by shortening the timing of diestrus and proestrus.

The observed changes in follicle development and estrous cycle of Abhd2^–/– females might not only result from an altered progesterone signaling but could also be influenced by a change in hormone synthesis. Ovarian hormone production is mainly governed by gonadotropins released by the pituitary gland. Although mRNA of Abhd2 has been detected in different brain regions, we have not been able to show the presence of the ABHD2 protein in areas other than the epithelial cells of the choroid plexus (Supplementary Figure 5), thus, making it unlikely that the observed ovarian phenotype would be caused by changes in neuronal function. To study the effect of Abhd2 ablation on steroidogenesis in the ovaries, qPCR was performed to detect expression levels of enzymes involved in hormone synthesis. Ovaries of superovulated Abhd2^–/– females showed no difference in expression of the follicle-stimulating hormone receptor (Fshr), a protein vital for follicle development and hormone synthesis, compared with ovaries of Abhd2^+/+ mice (Figure 6A). Neither was there a difference in the expression of cytochrome P450 family members (Cyp11a1, Cyp17a1, or Cyp19a1), the enzymes required for synthesis of progesterone, androgens, and estrogens, respectively. Since increased vascularization of the ovary precedes ovulation, the expression of vascular endothelial growth factor A (Vegfa), the factor required for vascular endothelial proliferation was also determined. However, no difference in expression between Abhd2^–/– and Abhd2^+/+ was detected (Figure 6A). The expression of nerve growth factor (Ngf) and its receptor [nerve growth factor receptor (Ngfr)] that are involved in ovulation and in the estrous cycle, were not different between Abhd2^–/– and Abhd^+/+ females. However, the expression of the NGF receptor neurotrophic receptor tyrosine kinase 1 [Ntrk1, previously known as tropomyosin receptor kinase A (TrkA)] was significantly increased in KO ovaries compared with those of wild-type mice (Figure 6B). It is interesting to note that the expression of NGF in stromal cells of the rodent ovary resembles that of ABHD2 (Dissen et al., 1996). Furthermore, increased NGF levels caused the mice to remain in the estrus stage even though they are still cycling (Dissen et al., 2009; Wilson et al., 2014). NGF signaling is also elevated in women with PCOS (Dissen et al., 2009; Gulino et al., 2016), while increased NGF signaling causes mice to show signs of PCOM, with a higher number of atretic antral follicles, without displaying cyst formation—the phenotype we also observed in Abhd2^–/– females.

The CRISPR-EZ method, where single guide RNA (sgRNA)/Cas9 complexes are delivered into mouse zygotes by electroporation, was developed by Chen et al. (2016) and Modzelewski et al. (2018).

SgRNAs to target the introns flanking Abhd2 exon 6 were designed as previously described. Briefly, we utilized the Gene Perturbation Platform (Doench et al., 2016), Chop-Chop (Montague et al., 2014), and CRISPR Design (Hsu et al., 2013) algorithms to design sgRNAs for the target sequence. The obtained 20-nt sequences were incorporated into a DNA oligonucleotide template containing a T7 promoter and a sgRNA scaffold by overlapping PCR using Phusion high fidelity DNA polymerase (New England Biolabs, M0530). The primer sequences for overlapping PCR were as previously described (Chen et al., 2016) including the designed sgRNA sequences; sgRNA1: 5′-TGTGCTGAGAAAACGCAGGT-3′ and sgRNA2: 5′-TGTCAACTGAAGGACCCAAG-3′) The template sequence was transcribed into RNA using a T7 RNA polymerase (New England Biolabs, E2040S) after which the DNA template was removed by treatment with RNase-Free DNaseI (New England Biolabs, M0303S). The produced sgRNAs were further purified by allowing them to bind to SeraMeg Speedbeads magnetic carboxylate-modified particles (GE Healthcare, 65152105050250). After washing the bead/RNA pellets with 80% ethanol the sgRNAs were eluted in nuclease-free water (Ambion, AM9937) and stored at −80°C until use.

To form a ribonucleoprotein complex, the Cas9 protein (QB3 Macrolab, University of California at Berkeley) was incubated with the two sgRNAs in a solution containing 20 mM HEPES (Sigma, H7523), pH 7.5, 150 mM KCl (Fisher Chemical, P217), 1 mM MgCl₂ (Sigma, 68475), 10% glycerol, (Sigma, G2025), and 1 mM reducing agent DL-dithiothreitol (Sigma, D0632), to a final molar ratio of 1:2. The ribonucleoprotein mixture was prepared at 37°C for 10 min immediately before electroporation. To obtain zygotes, 4-week-old C57Bl/6N (Charles River) female mice were superovulated by intraperitoneal injection (i.p.) of 5 IU (PMSG, Sigma, G4877), and 48 h later, 5 IU (hCG, Millipore, 230734), after which they were immediately put in mating with fertile 2- to 4-month-old C57Bl/6N males. Twelve hours post coitum, females with a plug were euthanized, and the one-cell zygotes were collected from the ampulla of the oviduct. The cumulus cells surrounding the fertilized eggs were removed by hyaluronidase treatment (Life Global Group, LGHY-010), and the zona pellucida was weakened by incubation in acidic Tyrode’s solution (Sigma, T1788) as previously described (Chen et al., 2016). For delivery of the sgRNA/Cas9 complex, ∼40 zygotes were pooled in Opti-MEM reduced serum media (Gibco, 31985-070) containing the preformed Cas9/sgRNA ribonucleoproteins and loaded into a 1-mm electroporation cuvette (BioRad, 165-2089). Electroporation was performed using the GenePulser Xcell (BioRad, 1652660) with six pulses at 30 V for 3 ms, separated by a 100-ms interval. Immediately after electroporation, the zygotes were washed in KSOM + AA media (KCl-enriched simplex optimization medium with amino acid supplement, Zenith Biotech, ZEKS-050) supplemented with 4 mg/ml bovine serum albumin (BSA, Sigma, A9647), and cultured overnight in KSOM/BSA at 37°C, 5% CO₂. The following day, the embryos that had developed into two-cell stage were implanted into pseudopregnant CD-1 females (Envigo) at ∼10 embryos per oviduct by the UC Berkeley Cancer Research Laboratory, Gene Targeting Facility. The resulting progeny was genotyped using EmeraldAmp GT PCR Master Mix (Takara, RR310) and primers flanking the deleted region of Abhd2, Abhd2 Fw: 5′-AGGGCTTAACTCTTGCTGGT-3′, Abhd2 Rev: 5′-ACTCAGACACGATCCGAGAC-3′, Tm: 56°C. Each mouse, positive for the deleted region of Abhd2, was placed in breeding with a C57Bl/6N mouse of the opposite gender. The heterozygous progeny of these matings were further bred to produce Abhd2^–/– mice and the control Abhd2^+/+ littermates. All mice were kept in the Animal Facility of the University of California, Berkeley, fed with standard chow diet (PicoLab Rodent diet 20, #5053, LabDiet) and hyper-chlorinated water ad libitum in a room with controlled light (14-h light, 10-h darkness) and temperature (23 ± 0.5°C).

To determine the breeding efficiency of Abhd2^+/+, Abhd2^+/–, and Abhd2^–/– female mice, 2-month-old females of each genotype were placed in breeding with either Abhd2^+/+ males (Abhd2^+/+ and Abhd2^–/– females) or Abhd2^+/– males (Abhd2^+/– and Abhd2^–/– females) for ∼6 months. Two or more breeding pairs of each genotype combination were used to determine the average number of offspring per litter and the birth rate of female mice. Furthermore, to detect the number of spontaneously ovulated eggs, 2- to 3-month-old females in proestrus were placed in breeding with fertile Abhd2^+/+ male mice. When a copulatory plug was detected, the females were euthanized, and the fertilized zygotes were collected from the oviduct ampulla in Dulbecco’s phosphate-buffered saline (DPBS, Gibco, 14190-144) and counted.

Sperm from the cauda epididymides of 2-month-old C57Bl/6N male mice were allowed to swim out in Embryomax Human Tubal Fluid medium (HTF, Millipore, MR-070-D) and incubated for capacitation, 1 h at 37°C, 5% CO₂. To collect eggs from seven Abhd2^+/+, eight Abhd2^+/–, and seven Abhd2^–/– females, superovulation of 3.5- to 4-week-old mice was performed using i.p. injection of PMSG and hCG as described above. Thirteen hours after hCG injection, the females were euthanized, and the cumulus–oocyte complexes were collected from the oviduct ampulla in HTF medium and counted. IVF was performed as previously described (Navarrete et al., 2015) using 300,000 sperm/ml HTF medium. After fertilization, the zygotes were washed in KSOM + AA media, supplemented with 4 μl of BSA and divided into ∼15 zygotes/10 μl of KSOM/BSA for culture at 37°C, 5% CO₂. Three and a half days following IVF, the number of fertilized eggs was determined as the percentage of embryos that had reached morula or blastula stage.

Each stage of the mouse estrous cycle was determined in the afternoon by cytological assessment of vaginal smear samples. Samples were collected from 2.5-month-old Abhd2^+/+, Abhd2^+/–, and Abhd2^–/– mice, three females of each genotype, every day for 1 month. DPBS was used to collect samples of each mouse, using the methods previously described (McLean et al., 2012). Samples were separately placed on glass slides and examined under an Olympus IX-75 inverted light microscope for cellular contents. Visual representation of each estrous cycle phase indicated by Zenclussen et al. (2014) was used as a reference.

To count the total number of follicles in Abhd2^+/+, Abhd2^+/–, and Abhd2^–/– ovaries, 3- to 4 3.5-month-old female mice of each genotype were euthanized at proestrus stage, and the ovaries were dissected out and weighed. The larger of the two ovaries was fixed in 4% paraformaldehyde (Electron Microscopy Sciences, 15714), embedded in optimal cutting temperature compound (OCT, Sakura, 4583), frozen on dry ice, and stored at −80°C until use. The ovaries were sectioned through, and every fifth section of 8 μm was mounted on glass slides and stained with hematoxylin 1 (Fisher HealthCare, 220-101) using standard procedures. Images of each ovarian section were taken at ×50 magnification and organized sequentially for follicle counting. Using this method, we were able to only count each follicle once, even though the larger follicles were present in several sections. Each follicle was categorized as early antral, antral, or CL based on their morphology. Early antral follicles were identified by the presence of segmented cavities between multiple layers of granulosa cells surrounding the oocyte. Antral follicles were identified by the presence of a large continuous antral cavity, and CL by the absence of oocytes. Atretic antral follicles were defined by a thinning of the granulosa cell layer, which displayed many pyknotic cells, and the additional lack of a clear cumulus cell layer surrounding the oocyte. The morphology of the atretic antral follicles were further confirmed by comparing hematoxylin and TUNEL stainings.

Ovaries were retrieved from 11 20- to 21-day-old wild-type (C57Bl/6N) and seven Abhd2^–/– female mice as described previously (Xu et al., 2006; Skory et al., 2015). Briefly, ovaries were placed in dissection media [L15 media (Gibco, 11415-064), 0.5% PenStrep (Fisher Scientific, 15140-122), and 1% fetal bovine serum (FBS, X&Y Cell Culture, FBS-500-HI)], and the bursal sac and any excess material were removed. Each ovary was halved and incubated in collagenase media [Minimum essential medium alpha (αMEM, Gibco, 12561-056), 0.5% PenStrep, 0.8% type I collagenase (Worthington, LS004194), and 1% DNase I (Fisher Scientific, EN0521)] at 37°C and 5% CO₂ for 40 min. Preantral follicles were mechanically isolated using 28-gauge insulin needles in dissection media. Only intact follicles that were 240 ± 10 μm in diameter were isolated and incubated at 37°C and 5% CO₂ in maintenance media (αMEM, 0.5% PenStrep, and 1% FBS) for 2 h before encapsulation. Follicles were encapsulated in 0.7% alginate and cultured as previously described (Xu et al., 2006; Skory et al., 2015), with slight modifications. The encapsulated follicles were placed individually in a 96-well plate (Fisher Scientific, 08-772-53) and cultured for 4 days in 100 μl of growth media [αMEM, 3 mg/ml of BSA, 1 mg/ml of bovine fetuin (Sigma, F3385), 0.01 IU/ml of recombinant human FSH (NHPP, Dr. Parlow), 5 μg/ml of insulin, 5 μg/ml of transferrin, and 5 ng/ml of selenium (Sigma, I1884)]. At day 2 of culture, half of the growth media were exchanged. After 4 days in culture, the alginate was removed by incubating the follicles in alginate lyase media [αMEM, 0.5% PenStrep, 1% FBS, and 1 mg/ml of alginate lyase (Sigma, A1603)] at 37°C and 5% CO₂ for 30 min. Thereafter, antral follicles larger than 330 ± 15 μm in diameter were washed in ovulation media [αMEM, 10% FBS, 5 ng/ml of EGF (Sigma, E4127), 1.5 IU/ml of hCG, 5 μg/ml of insulin, 5 μg/ml of transferrin, and 5 ng/ml of selenium] and placed individually in a 96-well plate containing 100 μl of ovulation media per well. After 16 h at 37°C and 5% CO₂, the percentage of successful ovulation was calculated as the number of follicles with a clear visual evidence of follicle rupture and oocyte extrusion compared with the total follicle number.

To detect ABHD2 protein in ovaries and different brain regions, 3.5- to 4-week-old Abhd2^+/+, Abhd2^+/–, and Abhd2^–/– superovulated mice and a 2.5-month-old Abhd2^+/+ female, respectively, were euthanized, the tissue dissected and snap frozen, and protein was isolated using standard procedures. The samples were analyzed by Western blotting, using a rabbit polyclonal anti-ABHD2 antibody (1:10,000 dilution, Proteintech Group, 14039-1-AP) and a peroxidase-conjugated anti-rabbit secondary antibody (1:15,000 dilution, Abcam, ab6721). To ensure equal sample loading, the membrane was stripped by incubation with 1 Min Plus Strip (GM Biosciences, GM6011) according to the instructions of the manufacturer. Thereafter, the membrane was re-hybridized with a mouse monoclonal anti-actin antibody (1:5,000 dilution, Abcam, ab3280-500) and a peroxidase-conjugated goat anti-mouse secondary antibody (1:15,000 dilution, EMD-Millipore, AP181P).

Ovaries of superovulated 3.5- to 4-week-old Abhd2^+/+ and Abhd2^–/– mice were fixed overnight at 4°C in 4% PFA, and the ovary of a 1-week-old Abhd2^+/+ female was fixed for 4 h after which they went through a sucrose gradient (10−20−30% sucrose/PBS) and were frozen in OCT. To detect localization of ABHD2 in the ovary, 8-μm sections were placed on charged slides and immediately used for staining, according to standard procedures. In brief, for antigen retrieval, the sections were incubated in 1% SDS solution for 5 min, blocked in 5% BSA for 1 h at room temperature, and incubated overnight at 4°C with rabbit polyclonal anti-ABHD2 (1:300) and mouse monoclonal anti-α-actin (1:500, sc-32251, Santa Cruz Biotechnology). The antibody–antigen complexes were visualized by incubation for 1 h at room temperature with 1:2,000 Alexa Fluor 488-conjugated goat anti-rabbit (Molecular Probes A11008/Jackson ImmunoResearch, 111-485-144) and Cy5-conjugated donkey anti-mouse (Jackson ImmunoResearch, 715-175-150) or anti-rabbit (Jackson ImmunoResearch, 711-015-152) antibodies. The sections were mounted using ProLong Gold antifade reagent with DAPI (Invitrogen, P36935) and imaged using confocal laser scanning microscopy (Olympus Fluoview FV1000).

For analysis of gene expression, 21- and 28-day-old Abhd2^+/+ mice were euthanized, and the ovaries were dissected out and snap frozen. Samples from superovulated 1- to 2-month-old mice at proestrus and estrus were similarly collected and frozen. Total RNA was isolated from the tissues using TRIzol reagent according to the instructions of the manufacturer (Ambion, 15596018). For reverse transcription, 1 μg of total RNA was treated with DNaseI and reverse-transcribed by using the RevertAid H Minus RT enzyme (Fisher Scientific, EP0451). The cDNA was diluted 1:50–1:100 for qPCR. qPCR was performed using the DyNAmo HS SYBR Green qPCR Master Mix (Fisher Scientific, F-410). All samples were run in triplicate reactions. For analysis of Abhd2 expression at different time points in wild-type mice ovaries primers binding to exon 3 and exon 4 were used (Abhd2 e3 Fw and Abhd2 e4 Re). To determine expression of the deleted exon 6 in Abhd2, KO mice primers binding in exon 5 and exon 6 were used (Abhd2 e5 Fw and Abhd2 e6 Re). Ribosomal protein L19 (Rpl19) and peptidylprolyl isomerase A (Ppia) were used as endogenous controls to equalize for the amounts of RNA in the ovaries. Primer sequences for Rpl19, Ppia, and cytochrome P450 family 19 subfamily A member 1 (Cyp19a1) were as previously described (Björkgren et al., 2012; Hakkarainen et al., 2015). Primer sequences and qPCR conditions for analyzing the expression of Abhd2, Fshr, cytochrome P450 family 11 subfamily A member 1 (Cyp11a1), cytochrome P450 family 17 subfamily A member 1 (Cyp17a1), Vegfa, Ngf, Ntrk1, and Ngfr are described in Supplementary Table 1.

To detect the effect of Abhd2 ablation on glucose uptake, 2.5-month-old Abhd2^+/+ (n = 5), Abhd2^+/– (n = 7), and Abhd2^–/– (n = 6) female mice were fasted overnight (16 h). Blood glucose levels were measured as mmol/L by tail vein sampling using Contour Next ONE meter (Ascensia Diabetes Care) and Contour Next blood glucose test strips (Ascensia Diabetes Care). Immediately after baseline measurement (0 min), the mice were given an i.p. injection of 2 g/kg glucose (20% solution), and blood glucose levels were measured at 15, 30, 60, and 120 min after injection.

For statistical analyses of fertility efficiency, ovulation number, days in estrus, follicle numbers, and gene expression levels, the GraphPad Prism 5 software (GraphPad Software, Inc., La Jolla, CA, United States) was used. Unpaired t-test was used to determine statistical significance, assigning p ≤ 0.05 as the limit. All results are shown with standard error of mean.