Proteomic quantification of native and ECM-enriched mouse ovaries reveals an age-dependent fibro-inflammatory signature

Detergent treatment effectively enriches the mouse ovarian ECM

To investigate age-associated changes to the ovarian proteome, we isolated ovaries from reproductively young (6–12 weeks) and old mice (10–12 months) and analyzed them both in their native state and enriched for the ECM to specifically determine how aging alters the ovarian matrix (Figure 1). We, therefore, first optimized an ECM enrichment strategy using a decellularization approach, which removes cellular and nuclear material while preserving the composition and structure of the ECM [30, 31]. Decellularized scaffolds are often used for bioengineering applications, as they can be re-seeded with allogenic cells, which has the benefit of preventing an immune response while maintaining tissue-specific cell phenotypes and functions [32, 33]. We further optimized this technique to enrich for the ECM of mouse ovaries prior to proteomic analysis. We first tested several detergents previously used for decellularizing murine, bovine, porcine, and human ovaries, including 0.5% sodium dodecyl sulfate (SDS), 0.1% SDS, and 1% Triton-X [34–37]. Decellularization has previously been performed in mouse ovaries with the goal of use in bioengineering applications, and as the decellularized tissue had to be completely devoid of cells, this inherently created the tradeoff of compromised ECM integrity [36, 38]. In contrast, our goal was to preserve as much of the mouse ovarian ECM as possible to allow for a comprehensive analysis of age-dependent alterations to the matrix. In our experimental setup, treatment with 1% Triton-X for 24 hours did not result in decellularization, whereas treatment with both 0.5% SDS for 10 hours and 0.1% SDS for 24 hours effectively removed cellular content as evidenced by lack of nuclei in Hematoxylin and Eosin (H&E) stained tissue sections (Supplementary Figure 1). We evaluated matrix integrity using the histological stain PicroSirius Red (PSR) which detects collagen I and III fibers. Compared to 0.5% SDS, treatment with 0.1% SDS appeared to better preserve the integrity of the collagen-matrix (Supplementary Figure 1). Therefore, we further refined the ECM enrichment protocol using 0.1% SDS.

As longer exposure to detergent may compromise the ECM, we performed a time-course with 0.1% SDS treatment for 0, 10, 12.5, or 15 hours to establish the minimum amount of time needed to remove cells without perturbing ECM integrity. Although nuclear material was still visible following 10-hour treatment, most was effectively removed at 12.5 and 15 hours as assessed by H&E, Hematoxylin, and Hoechst staining of tissue sections to visualize DNA (Figure 2A–2C). However, the structure of the collagen-matrix as assessed by PSR staining was better maintained following 0.1% SDS treatment for 12.5 hours compared to 15 hours (Figure 2D). In fact, ovarian structures, including corpora lutea, were still distinguishable in tissue sections from ovaries treated with 0.1% SDS for 12.5 hours (Figure 2A). Thus, we used 0.1% SDS for 12.5 hours to enrich for the ovarian ECM for all downstream studies.

ECM-enrichment was effective in ovaries from reproductively young and old mice

To validate that our optimized ECM enrichment strategy was equivalently effective irrespective of animal age, we incubated ovaries from reproductively young and old mice in 0.1% SDS for 12.5 hours in parallel and evaluated ovarian architecture and cellular content through assessment of DNA using various approaches. These conditions maintained the overall structure of the ovaries with regions previously occupied by follicles, corpora lutea, and stroma still clearly visible (Figure 3A). The majority of the H&E-stained tissue lacked visible nuclei, with 95.3 ± 3.1% and 89.7 ± 3.2% of the ovaries from reproductively young and old mice, respectively, being enriched in decellularized matrix (Figure 3A, 3B). We further confirmed these findings by demonstrating the absence of nuclear material in tissue sections from ECM-enriched ovaries relative to native ovaries from reproductively young and old mice stained with Hematoxylin only or Hoechst to detect DNA (Figure 3C, 3D). Moreover, we validated these histological findings by running DNA extracted from native and ECM-enriched ovaries on agarose gels. Whereas DNA bands were clearly detectable in native ovaries, this was not the case in detergent-treated samples irrespective of animal age, further demonstrating effective removal of cellular contents (Figure 3E, 3F). We optimized conditions such that a small percentage of residual nuclei in addition to identifiable ovarian structures were maintained in ECM-enriched tissues, thus maximizing the likelihood that the detergent treatment did not completely strip the tissue to the point of perturbing ECM integrity. Our detergent treatment approach also maintained the integrity of the ovarian ECM as evidenced by positive collagen I and III signal in ECM-enriched ovaries from both reproductively young and old mice, which was similar to that observed in native ovaries (Figure 4). With advanced reproductive age, there is a reported accumulation of collagen I and III in the ovary which ultimately contributes to tissue fibrosis [13, 19, 20]. Although the reproductively old mice used in this study are not as old as those in previous studies where significant age-associated fibrosis was observed, we did see a similar trend of increased collagen in our samples. For example, there was a 2.3 ± 1.0-fold increase in PSR-positive area in native ovaries from reproductively old mice compared to young counterparts (Figure 4A, 4B). In the ECM enriched samples, there was a 1.1 ± 0.2-fold increase in PSR-positive area with age (Figure 4C, 4D). Thus, our ECM enrichment strategy preserved key matrix components and architecture.

Native and ECM-enriched ovaries have distinct age-associated proteomic signatures

To determine how reproductive aging impacts the ovarian proteome and ECM, we used an unbiased proteomic approach using data-independent acquisition on an Orbitrap Exploris 480 and a direct-DIA method in which the DIA-MS data were used to simultaneously identify and quantify all proteins (Figure 1). For each of the 5 biological replicates per age group, one ovary from each mouse was analyzed in its native state, while the contralateral ovary was enriched for the ECM. Using this experimental approach, we probed age-dependent changes to the whole ovarian proteome and ECM in parallel (Figure 1). For the label-free quantitative proteomic workflow, we lysed the native tissue with 2% SDS and resolubilized the ECM with 2% SDS and 50 mM TEAB, respectively. Samples were then proteolytically digested with trypsin using S-Trap micro spin columns. Following DIA-MS analysis, all data were searched and statistically processed by Spectronaut v14, and subjected to additional pathway analysis and data mining (Figure 1).

Partial Least Squares-Discriminant Analysis (PLS-DA) revealed separated clustering of both native and ECM-enriched samples based on age, indicating that ovaries from reproductively young and old mice have distinct proteomic signatures (Figure 5A, 5B). Proteomic analysis identified 4,084 proteins in native mouse ovaries and 3,160 proteins in ECM-enriched mouse ovaries, of which 2,523 were present in both native and ECM-enriched samples (Supplementary Tables 1 and 2; Supplementary Figure 2). ECM-enriched ovaries had a different proteome from native counterparts, which in fact, included 637 proteins not detected in native samples (Supplementary Table 2; Supplementary Figure 2). The difference in the number of proteins identified between native and ECM-enriched ovaries was as anticipated, given the removal of cellular proteins in the ECM-enriched samples. All proteins in native and ECM-enriched samples were identified if ≥2 unique peptides per protein were detected in both reproductively young and reproductively old ovary samples. These proteins were subsequently quantified across all conditions using our data-independent acquisition approach and generating relative quantification results. We did not quantify changes between native and ECM-enriched samples as this was not the focus of our study.

Overall, 180 proteins were significantly altered (q-value < 0.01 and absolute average log₂ ratio > 0.58) with advanced reproductive age in native ovaries (Supplementary Table 3), and 279 proteins were significantly altered with age in ECM-enriched ovaries (Supplementary Table 4), of which 76 proteins were common between groups (Supplementary Figure 2). The most highly upregulated proteins with advanced reproductive age included immunoglobulins, ECM proteins such as tenascin-C, tenascin-X, and TGFβ-induced protein, proteins that regulate metal homeostasis such as ferritin and ceruloplasmin, as well as tumor suppressors such as SDH enzyme and glycoprotein NMB (Figure 5C, 5D). Significantly downregulated proteins with age included oocyte-expressed protein homolog (OOEP) as well as NOD-, LRR-, and pyrin domain-containing proteins 5 and 14 and KH domain-containing protein 3 (Figure 5C, 5D). These proteins are expressed in the oocyte or are required for early embryonic development, and thus they are anticipated to decrease given the age-dependent loss of germ cells. The identification of oocyte-specific proteins, including OOEP, in ECM-enriched ovaries was surprising given the several wash steps with large volumes and agitation that follow detergent treatment in our decellularization protocol, which we would expect to remove most proteins from lysed cells. However, although the majority of H&E-stained stained tissue sections from ECM-enriched ovaries lacked visible nuclei, there were still some cells remaining, which may explain the detection of these cellular proteins. These conditions for decellularization were selected intentionally to enrich for the ECM, while still preserving the integrity of the matrix, which resulted in the tradeoff of some residual cells.

Core matrisome and matrisome-associated proteins are significantly altered with age

To annotate core matrisome (collagens, proteoglycans, and ECM glycoproteins) and matrisome-associated proteins (ECM regulators, ECM-affiliated proteins, and secreted factors) in the mouse ovary, we utilized Matrisome DB, a database that compiles in silico and experimental data on matrisome proteins [39, 40]. We identified 81 core matrisome and 93 matrisome-associated proteins in native ovaries, as well as 74 core matrisome and 61 matrisome-associated proteins in ECM-enriched ovaries (Supplementary Table 5). Surprisingly, only seven matrisome proteins were specific to ECM-enriched samples, whereas 45 matrisome proteins were specific to native samples (Supplementary Table 5). Five secreted factors (anti-mullerian hormone, S100 calcium-binding protein A1, hepatocyte growth factor activator, secreted frizzled-related protein 1, and follistatin) were among the proteins specific to native ovaries (Supplementary Table 5). Growth factors are often sequestered in the ECM and the detection of some secreted factors in native ovaries that were not identified in ECM-enriched samples was anticipated given that detergent treatment and subsequent wash steps may remove some of the loosely bound proteins. However, eleven growth factors were identified in both native and ECM-enriched samples, further supporting that our conditions for ECM-enrichment did not perturb overall matrix integrity (Supplementary Table 5).

Of the proteins that were upregulated with advanced reproductive age in either native or ECM-enriched samples or in both groups, 15 were core matrisome proteins and 22 were matrisome-associated proteins (Figure 6A: Supplementary Table 6). Eleven of the proteins downregulated in the ovary with advanced reproductive age were part of the core matrisome and ten were matrisome-associated proteins (Figure 6B; Supplementary Table 5). Overall, 14 of the core matrisome and matrisome-associated proteins upregulated with age were specific to the native ovary, 11 were specific to the ECM-enriched ovary, and 12 were present in both (Supplementary Table 6). Most of the core matrisome and matrisome-associated proteins downregulated with age were specific to the ECM-enriched ovary (13 proteins), with only four downregulated proteins specific to the native ovary and four proteins present in both samples (Supplementary Table 6). Of the 13 proteins downregulated with age specific to the ECM-enriched ovary, six of them were ECM glycoproteins including thrombospondin-1, fibrillin-2, and three zona pellucida proteins, which are specialized glycoproteins that surround the oocyte (Supplementary Table 6). Ovarian collagen, collagen VI α6, was only detected in the native ovary and was upregulated with age, whereas collagen XI α1 was distinctly detected in the ECM-enriched ovary and was downregulated with age (Supplementary Table 6). Overall, these data demonstrate that ECM proteins are some of the most robustly dysregulated in the ovary with advanced reproductive age.

To determine if the proteins upregulated with age in the mouse ovary may be implicated in age-associated ovarian fibrosis, we cross-referenced our dataset with FibroAtlas, a database of genes associated with fibrosis in several organ systems [41]. In fact, 17 of the matrisome proteins upregulated with age in the native ovary and 13 of the matrisome proteins upregulated with age in the ECM-enriched ovary have been previously associated with fibrosis in other organs (Table 1).

Pathways associated with genomic stability and proteostasis are downregulated in the ovary with age

Gene ontology analysis of proteins significantly altered with advanced reproductive age revealed that biological processes associated with genomic stability and epigenetic regulation, including “Nuclear DNA replication”, “Nucleosome organization”, “Chromatin assembly”, “DNA metabolic process”, and “Chromosome organization” were downregulated with age in native ovaries (Figure 7A; Table 2). Interestingly, several members of the mini-chromosome maintenance (MCM) family of DNA replication licensing factor proteins (MCM2, MCM3, MCM4, MCM6, and MCM7) drove these GO terms (Table 2). DNA replication, epigenetic modification, and DNA packaging-related molecular functions were also downregulated in both native and ECM-enriched ovaries with age (Supplementary Figure 3). GO terms associated with DNA helicase activity, DNA methyltransferase activity, and histone binding were downregulated in the ovary with age (Supplementary Figure 3). Moreover, biological processes associated with maintaining proteostasis, including “Cellular amino acid biosynthetic process” and “Ribosome biogenesis” were downregulated with age in both native and ECM-enriched ovaries (Figure 7A, 7B).

Pathways associated with inflammation and ECM organization are upregulated in the ovary with age

Biological processes associated with immune response were enriched in gene ontology analysis of proteins significantly altered with age in native and ECM-enriched ovaries (Figure 8A, 8B; Table 3). These upregulated immune-related pathways included “Antibacterial humoral response”, “Complement activation”, “Humoral response mediated by circulating immunoglobulins”, “Lymphocyte mediated immunity”, and “Regulation of immune response” (Figure 8A, 8B; Table 3). Age-associated upregulation of “Complement activation” is interesting given the complement system has been implicated in several age-related diseases, including Alzheimer’s disease and age-related macular degeneration [42]. In the ovary, however, the majority of research on the complement cascade has been done in the context of ovarian cancer [43]. Several immunoglobulins, complement component 4b (C4B), complement component receptor 1-like protein (CR1L), and the mitochondrial protein complement C1q binding protein (C1QBP) are among proteins that drive this GO term (Table 3).

Increased inflammation in the aging ovary may be in part due to increased oxidative stress and consistent with this, molecular functions associated with oxidoreductase activity were upregulated with age (Supplementary Figure 4) [44–46]. Chronic inflammation in the aging ovary has been associated with the presence of a unique population of multinucleated macrophage giant cells (MNGCs) [13, 47]. Although these ovarian MNGCs have not been well characterized, osteoclasts are one of three main subtypes of MNGCs [48]. Thus, upregulation of “Chondrocyte differentiation”, “Osteoblast differentiation”, and “Bone development” in the native ovary with age may be indicative of MNGC signaling and proteins that underlie these GO terms including transmembrane glycoprotein NMB (GPNMB) and transforming growth factor-β-induced protein (TGFBI) may be putative markers for ovarian MNGCs (Figure 8A; Table 4). Macrophage fusion to form MNGCs allows for enhanced phagocytosis and the “Membrane invagination” biological process was upregulated in the native ovary with age, consistent with this potential increase in phagocytic activity (Figure 8A; Table 4) [49].

Inflammation often precedes fibrosis, and biological processes relevant to ECM remodeling and fibrosis were also upregulated with age, including “Extracellular matrix organization” and “Wound healing” in native ovaries, as well as “Cellular response to fibroblast growth factor stimulus” in both native and ECM-enriched ovaries (Figure 8A; Table 5). Although age-associated ovarian fibrosis has been well characterized, the mechanisms that underlie this fibrosis are not well understood and further research is required to determine how proteins including tenascin-C (TNC), cathepsin family members, and transforming growth factor-β-induced protein (TGFBI), which drive fibrosis-related GO terms may contribute to this robust ovarian aging phenotype (Table 5). Binding of core matrisome components and matrisome-associated proteins, including syndecans, proteoglycans, collagens, and glycosaminoglycans, were among molecular functions upregulated with age in both native and ECM-enriched ovaries (Supplementary Figure 4). Age-associated fibrosis in the ovary contributes to increased tissue stiffness, and consistent with this, the “Extracellular matrix constituent conferring elasticity” pathway was downregulated in native ovaries from reproductively old mice (Supplementary Figure 3) [19, 21].

Animals

All experiments were performed using female CD-1 mice purchased from Envigo (Indianapolis, IN, USA) except for a small subset of optimization experiments which were performed with female CB6F1 purchased from Envigo (Supplementary Figure 1). Reproductively young mice were used for experiments at 6–12 weeks and reproductively old mice were used at 10–12 months of age. Reproductively young mice were virgin animals, whereas our reproductively old mice were retired breeders due to the resources required to age animals for this amount of time. At 10–12 months of age mice display reproductive aging phenotypes, including subfertility, fibrotic foci in the ovarian stroma, reduced follicle quantity, and decreased chromosome cohesion [13, 85–87]. Mice were housed in a controlled barrier facility at Northwestern University’s Center for Comparative Medicine (Chicago, IL, USA) under constant temperature, humidity, and light (14 h light/10 h dark). Upon arrival to Northwestern University, animals were provided water and Teklad Global irradiated 2016 chow (Envigo, Madison, WI, USA) containing minimal phytoestrogens, ad libitum. All mice were allowed to acclimate for a minimum of one week upon arrival in our vivarium prior to experimental use. All experiments were performed under protocols approved by the Institutional Animal Care and Use Committee (Northwestern University) and in accordance with the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals.

ECM enrichment

To enrich for the ECM, ovaries were washed in Dulbecco’s Phosphate-Buffered Saline (DPBS, Gibco, Grand Island, NY, USA) and immersed in DPBS containing 0.1% (w/v) sodium dodecyl sulfate (SDS, Sigma-Aldrich, St. Louis, MO, USA) for 12.5 hours, or specific times as described in the figure legends, at room temperature on a nutating rocker. Ovaries were then washed in 15 mL RO water (4 × 30 minutes) and 1 mL DPBS (1 × 1 hour) and incubated in DPBS containing 40 units/mL Deoxyribonuclease I (Sigma-Aldrich, St. Louis, MO, USA) for 30 min at 37°C. Ovaries were subsequently washed in 15 mL DPBS (2 × 30 minutes) and flash-frozen on dry ice or fixed in Modified Davidson’s (Electron Microscopy Sciences, Hatfield, PA, USA) overnight at 4°C. ECM enrichment in Supplementary Figure 1 was performed in DPBS containing 0.5 or 0.1% (w/v) SDS or 1% Triton X-100 (TX-100, Alfa Aesar, Haverhill, MA, USA) following the same procedure. Native ovaries were washed in DPBS and flash-frozen on dry ice or fixed in Modified Davidson’s overnight at 4°C immediately after harvesting.

To validate ECM-enrichment by measuring residual double stranded DNA, flash-frozen ovaries were lyophilized (Freezone 6, Labconco, Kansas City, MO, USA) at the Northwestern University Analytical bioNanoTechnology Core Facility (ANTEC). DNA was extracted from lyophilized ovaries using a Quick-DNA MidiPrep Plus Kit (Zymo Research, Irvine, CA, USA) following the manufacturer’s instructions. Following extraction, DNA was run on a 1% (w/v) agarose gel with a 1 kb DNA ladder (New England Biolabs, Ipswich, MA, USA). Agarose gels were imaged using a Bio-Rad ChemiDoc Imaging System (Bio-Rad, Hercules, CA, USA).

Tissue processing and histochemical staining

Following fixation, native and ECM-enriched ovaries were transferred to 70% ethanol and stored at 4°C until processing. Samples were dehydrated using an automated tissue processor (Leica Biosystems, Buffalo Grove, IL, USA), embedded in paraffin, and sectioned (5 μm thickness) using a Reichert-Jung Biocut 2035 microtome (Leica Biosystems, Buffalo Grove, IL, USA).

Hematoxylin and eosin (H&E) staining was performed using a Leica Autostainer XL (Leica Biosystems, Buffalo Grove, IL, USA). Tissue sections were cleared with Xylene (Mercedes Scientific, Lakewood Ranch, FL, USA) in 3, 5-minute incubations and mounted with Cytoseal XYL (ThermoFisher Scientific, Waltham, MA, USA). Hematoxylin staining was performed following a standard protocol. Tissue sections were cleared with Citrosolv (Fisher Scientific Pittsburgh, PA, USA) in 3, 5-minute incubations and mounted with Cytoseal XYL.

Picrosirius Red (PSR) staining was performed as previously published [13, 19]. Briefly, tissue sections were deparaffinized in Citrosolv, rehydrated in graded ethanol baths (100, 70, and 30%) and washed in RO water. Slides were immersed in PSR staining solution for 40 minutes, then incubated in acidified water (0.05 M hydrochloric acid) for 90 seconds. Tissue sections were dehydrated in 100% ethanol, cleared in Citrosolv, and mounted with Cytoseal XYL.

For Hoechst staining, tissue sections were deparaffinized in Citrosolv (2, 3-minute incubations), rehydrated in graded ethanol baths (100, and 95%) and washed in RO water and PBS. Slides were incubated with Hoechst (NucBlue, Invitrogen, Carlsbad, CA, USA; 2 drops per mL PBS) for 30 minutes in a humidified chamber. Slides were washed in PBS and mounted in Vectashield antifade mounting medium (Vector Laboratories, Burlingame, CA, USA).

Imaging and image analysis

Brightfield images were taken with an EVOS FL Auto Cell Imaging System (ThermoFisher Scientific, Waltham, MA, USA) using a 10×, 20×, or 40× objective. To view entire ovarian tissue sections, scans comprised of a series of individual images were taken across the tissue and then automatically stitched together using the EVOS software. Epifluorescence images were taken with an EVOS FL Auto Cell Imaging System equipped with a DAPI LED light cube (Excitation 357/44 nm, Emission 447/60 nm) using a 10× or 40× objective. Imaging settings, including light, gain, and exposure times, were kept consistent between samples.

Image processing and analysis was performed using FIJI software (National Institutes of Health, Bethesda, MD). Relative area of PSR-positive signal was quantified above a threshold that was set based on the section with the most staining, as previously published [13, 19]. This threshold was kept constant for all images analyzed for each experiment and fold-change was calculated over young ovarian sections. To quantify relative ECM-Enriched area from Hematoxylin-stained tissue sections, thresholding was used to measure total ovarian area and area with residual nuclei was measured manually using the freehand tool in FIJI.

Statistical analysis

Statistical analysis was performed using GraphPad Prism Software (La Jolla, CA, USA). T-tests were performed to evaluate differences between groups. P values < 0.05 were considered significant. The number of biological replicates (N) for each experiment is listed in the figure legends.

Tissue lysis and homogenization

Ovarian tissue from reproductively young (N = 5; 5 native and 5 ECM-enriched ovaries) and reproductively old (N = 5; 5 native and 5 ECM-enriched ovaries) mice was prepared for proteomic analysis. Samples were homogenized in 600 μL lysis buffer containing 8 M urea, 2% sodium dodecyl sulfate (SDS), 1 μM trichostatin A (TSA), 3 mM nicotinamide adenine dinucleotide (NAD), 75 mM sodium chloride, and 1× protease and phosphatase inhibitor cocktail (Thermo Fisher Scientific, Waltham, MA, USA) in 200 mM triethylammonium bicarbonate (TEAB) by adding them to 2.0 mL safe-lock tubes (VWR International, Radnor, PA, USA) containing stainless steel beads and subjected to three intervals of high-speed shaking (25 Hz, 1 min) using a Qiagen TissueLyser II (Qiagen, Hilden, Germany). Tissue homogenates were centrifuged at 15,700 × g for 10 min at 4°C, and the supernatant was collected for label-free quantitative proteomics experiments. Protein concentration was determined using Bicinchoninic Acid (BCA) assay (Thermo Fisher Scientific, Waltham, MA, USA).

Protein digestion and desalting

Aliquots of 200 μg protein lysates for each sample were brought to the same overall volume of 100 μL with water, reduced using 20 mM dithiothreitol in 50 mM TEAB at 50°C for 10 min, cooled to room temperature (RT) and held at RT for 10 min, and alkylated using 40 mM iodoacetamide in 50 mM TEAB at RT in the dark for 30 min. Samples were acidified with 12% phosphoric acid to obtain a final concentration of 1.2% phosphoric acid. S-Trap buffer consisting of 90% methanol in 100 mM TEAB at pH ~7.1, was added and samples were loaded onto the S-Trap micro spin columns (Protifi, Fairport, NY, USA). The entire sample volume was spun through the S-Trap micro spin columns at 4,000 × g at RT, binding the proteins to the micro spin columns. Subsequently, S-Trap micro spin columns were washed twice with S-Trap buffer at 4,000 × g at RT and placed into clean elution tubes. Samples were incubated for one-hour at 47^oC with sequencing grade trypsin (Promega, San Luis Obispo, CA) dissolved in 50 mM TEAB at a 1:25 (w/w) enzyme:protein ratio, and then digested overnight at 37^oC.

Peptides were sequentially eluted from S-Trap micro spin columns with 50 mM TEAB, 0.5% formic acid (FA) in water, and 50% acetonitrile (ACN) in 0.5% FA. After centrifugal evaporation, samples were resuspended in 0.2% FA in water and desalted with Oasis 10-mg Sorbent Cartridges (Waters, Milford, MA, USA). The desalted elutions were then subjected to an additional round of centrifugal evaporation and re-suspended in 0.1% FA in water at a final concentration of 1 μg/μL. Eight microliters of each sample was diluted with 2% ACN in 0.1% FA to obtain a concentration of 400 ng/μL. One microliter of indexed Retention Time Standard (iRT, Biognosys, Schlieren, Switzerland) was added to each sample, thus bringing up the total volume to 20 μL [88].

Mass spectrometric analysis

Reverse-phase HPLC-MS/MS analyses were performed on a Dionex UltiMate 3000 system coupled online to an Orbitrap Exploris 480 mass spectrometer (Thermo Fisher Scientific, Bremen, Germany). The solvent system consisted of 2% ACN, 0.1% FA in water (solvent A) and 80% ACN, 0.1% FA in ACN (solvent B). Digested peptides (400 ng) were loaded onto an Acclaim PepMap 100 C₁₈ trap column (0.1 × 20 mm, 5 μm particle size; Thermo Fisher Scientific) over 5 min at 5 μL/min with 100% solvent A. Peptides (400 ng) were eluted on an Acclaim PepMap 100 C₁₈ analytical column (75 μm × 50 cm, 3 μm particle size; Thermo Fisher Scientific) at 300 nL/min using the following gradient: linear from 2.5% to 24.5% of solvent B in 125 min, linear from 24.5% to 39.2% of solvent B in 40 min, up to 98% of solvent B in 1 min, and back to 2.5% of solvent B in 1 min. The column was re-equilibrated for 30 min with 2.5% of solvent B, and the total gradient length was 210 min. Each sample was acquired in data-independent acquisition (DIA) mode [27–29]. Full MS spectra were collected at 120,000 resolution (Automatic Gain Control (AGC) target: 3e6 ions, maximum injection time: 60 ms, 350-1,650 m/z), and MS2 spectra at 30,000 resolution (AGC target: 3e6 ions, maximum injection time: Auto, Normalized Collision Energy (NCE): 30, fixed first mass 200 m/z). The isolation scheme consisted of 26 variable windows covering the 350-1,650 m/z range with an overlap of 1 m/z [28].

DIA data processing and statistical analysis

DIA data were processed in Spectronaut (version 14.10.201222.47784) using directDIA. Data extraction parameters were set as dynamic and non-linear iRT calibration with precision iRT was selected. Data were searched against the Mus musculus reference proteome with 58,430 entries (UniProtKB-TrEMBL), accessed on 01/31/2018. Trypsin/P was set as the digestion enzyme and two missed cleavages were allowed. Cysteine carbamidomethylation was set as a fixed modification while methionine oxidation and protein N-terminus acetylation were set as dynamic modifications. Identification was performed using 1% precursor and protein q-value. Quantification was based on the peak areas of extracted ion chromatograms (XICs) of 3 – 6 MS2 fragment ions, specifically b- and y-ions, with local normalization and q-value sparse data filtering applied (Supplementary Tables 1, 2). In addition, iRT profiling was selected. Differential protein expression analysis comparing either (1) Young to Old Native or (2) Young to Old ECM-enriched was performed using a paired t-test, and p-values were corrected for multiple testing, using the Storey method [89]. Specifically, group wise testing corrections were applied to obtain q-values. Protein groups with at least two unique peptides, q-value < 0.01, and absolute Log₂(fold-change) > 0.58 were considered significantly altered (Supplementary Figure 2, Supplementary Tables 3, 4).

Statistical processing and pathway analysis

Partial least square-discriminant analysis (PLS-DA) of the proteomics data were performed using the package mixOmics in R (version 4.0.2; RStudio, version 1.3.1093) [90]. Volcano plots for differential analysis were generated using R (RStudio, version 1.3.1093). Annotation of matrisome proteins was performed using matrisome DB database [39, 40].

An over-representation analysis (ORA) was performed using Consensus Path DB-mouse (Release MM11, 14.10.2021), developed by the bioinformatics group at the Max Planck Institute for Molecular Genetics (Berlin, Germany) [91, 92]. The following comparisons were used to evaluate which gene ontology terms, including biological processes and molecular functions, were significantly enriched: (1) Old vs. Young Native and (2) Old vs. Young ECM-enriched. Gene ontology terms identified from the ORA were subjected to the following filters: q-value < 0.05, term category = b (biological processes) or m (molecular functions), and term level = 4 for biological processes or 3 for molecular functions. Dot plots were generated using the ggplot2 package in R (version 4.0.5; RStudio, version 1.4.1106) to visualize significantly enriched biological processes from each comparison [93].

Data availability

Raw data and complete MS data sets have been uploaded to the Mass Spectrometry Interactive Virtual Environment (MassIVE) repository, developed by the Center for Computational Mass Spectrometry at the University of California San Diego, and can be downloaded using the following link: https://massive.ucsd.edu/ProteoSAFe/dataset.jsp?task=13fe684e74b34dcaa087c6960e5a40ac (MassIVE ID number: MSV000091790; ProteomeXchange ID: PDX041789).