tissue (ICRS grade ≥ – ≥ 5; ICRS grade ≥ ≥ 6. Shaded boxes ≥ **Figure 2.** Gene expression levels of HBP-related enzymes 7 days after trauma and in highly degenerated tissue. Gene expression of HBP-related enzymes was evaluated in (**A**,**B**,**D**), impacted and subsequently treated cartilage explants, as well as (**E**) highly degenerated cartilage tissue (ICRS grade ≥ 3). Moreover, the ratio of OGA to OGT was calculated for (**C**) impacted and subsequent treated cartilage explants, as well as (**F**) OA cartilage. Statistical analysis: (**A**,**B**,**D**) one-way ANOVA, (**C**) the Kruskal–Wallis test, (**E**) multiple *t*-tests, (**F**) unpaired two-tailed *t*-test. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05; [versus C] c = *p* < 0.05, cc = *p* < 0.01; all data sets *n* ≥ 5; ICRS grade ≥ 3 tissue *n* ≥ 6. Shaded boxes = traumatized cartilage explants (T), striped boxes = highly degenerated cartilage tissue (ICRS grade ≥ 3).

−1 −2

≥ 5, except for 5 mM and 10 mM GS, **Figure 3.** Effects of glucosamine treatment or addition of inhibitors on cell viability 7 days after cartilage trauma. Impacted cartilage explants were treated with different concentrations of (**A**) glucosamine derivatives GlcNAc or GS or (**B**) OGA-inhibitor PUGNAc and GFAT-1-Inhibitor AZA, respectively, for 7 days. (**C**) Exemplary fluorescence images of live/dead staining. Statistical analysis: (**A**,**B**) one-way ANOVA. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05, \*\* = *p* < 0.01; [versus C] cc = *p* < 0.01; all data sets *n* ≥ 5, except for 5 mM and 10 mM GS, *n* = 3. Blank box = unimpacted control (C), shaded boxes = traumatized cartilage explants (T).

−2

NAc: − −3

–

– ≥ 5. **Figure 4.** Effects of glucosamine, PUGNAc, and AZA on chondroanabolic processes 7 days after cartilage trauma. Chondroanabolism was evaluated by means of gene expression analysis of (**A**) COL2A1, (**B**) HAS2, and (**C**) ACAN, as well as by (**D**) quantification of CPII release. Statistical analysis: (**A**,**B**) Kruskal–Wallis test, (**C**,**D**) one-way ANOVA. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05; [versus C] c = *p* < 0.05, cc = *p* < 0.01; all data sets *n* ≥ 5. Blank box = unimpacted control (C), shaded boxes = traumatized cartilage explants (T).

Compared to the unimpacted cartilage, chondroanabolic gene expression was significantly reduced after trauma (Figure 4A–C). Although, treatment with 2.5 mM GlcNAc enhanced the gene expression of COL2A1 ([vs. T] 2.4-fold) and ACAN ([vs. T] 1.6-fold), both glucosamine derivatives suppressed the biosynthesis of type II collagen ([vs. C] GlcNAc: −3.5 ng/mL; GS: −3 ng/mL; Figure 4D). Moreover, 1 mM GS further reduced gene expression of ACAN.

While AZA had rather anti-anabolic effects, as demonstrated by reduced biosynthesis of type II collagen ([vs. C] −2.3 ng/mL), PUGNAc significantly induced both gene expression of HAS2 ([vs. T] 2.4-fold) and the release of CPII ([vs. T] +2.2 ng/mL), despite unchanged mRNA levels of COL2A1.

#### *2.4. Trauma-Induced Expression of MMPs and Subsequent Type II Collagen Degradation Are Markedly Decreased after Treatment with Glucosamines or PUGNAc*

Possible influence of the HBP on trauma-induced expression of MMPs and subsequent breakdown of type II collagen was evaluated by gene expression analysis of MMP-1 and -13 (Figure 5A,B), quantification of secreted MMP-2 (zymographical detection; Figure 6 and MMP-13 (Figure 5C), as well as the degradation product of type II collagen (C2C), generated by MMP-1, -8, and -13 (Figure 5D).

T] GlcNAc: − −1

−0

T] − −0

**Figure 5.** Effects of glucosamine treatment or addition of inhibitors on trauma-induced expression of collagenases and COL2A breakdown 7 days after trauma. Catabolic processes were assessed by the means of gene expression analysis of (**A**) MMP-1 and (**B**) MMP-13, as well as (**C**) quantification of MMP-13 release and (**D**) type II collagen breakdown product C2C. Statistical analysis: (**A**,**B**) Kruskal–Wallis test, (**C**,**D**) one-way ANOVA. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05; [versus C] c = *p* < 0.05, cc = *p* < 0.01; all data sets *n* ≥ 5. Blank box = unimpacted control (C), shaded boxes = traumatized cartilage explants (T).

Cartilage trauma resulted in excessive gene expression of ECM-degrading MMPs ([vs. C] MMP1: 4-fold; MMP13: 5.8-fold), though concentration of C2C was not significantly enhanced compared to the unimpacted control. Both glucosamine derivatives exhibited anti-catabolic effects to varying degrees. While GlcNAc had stronger suppressive effects, with respect to the gene expression of MMP-1 (Figure 5A), GS revealed higher efficacy in the case of MMP-13 (Figure 5B). However, GlcNAc and GS demonstrated equal anti-catabolic and chondroprotective effects, respectively, concerning the secretion of MMP-13 ([vs. T] GlcNAc: −11.54 ng/mL; GS: −11.8 ng/mL; Figure 5C) and subsequent degradation of type II collagen ([vs. T] both −0.97 ng/mL; Figure 5D). Moreover, GlcNAc significantly reduced the conversion of pro-MMP-2 to active MMP-2 by 2.9-fold (Figure 6C).

Although, comparable anti-catabolic effects were found after inhibition of OGA by means of PUGNAc, as demonstrated by decreased secretion of MMP-13 ([vs. T] −11 ng/mL), the reduction of the degradation product C2C was less pronounced ([vs. T] −0.57 ng/mL). While no significant changes could be observed for the secretion of pro-MMP2 (Figure 6A), the addition of AZA increased the zymographically detectable amount of active MMP-2 ([vs. C] 4-fold; [vs. T] 3-fold; Figure 6B) and the generation of C2C ([vs. T] +0.5 ng/mL) after cartilage trauma.

– ≥ 5. Blank box **Figure 6.** Effects of glucosamine treatment or addition of inhibitors on trauma-induced secretion and activation of MMP-2 7 days after trauma. Release of (**A**) pro-MMP-2 and (**B**) active MMP-2 was measured by means of gelatin zymography, as exemplarily demonstrated in (**D**,**E**). (**C**) Ratio of zymographically detectable pro-MMP-2 to active MMP-2. Statistical analysis: (**A**–**C**) one-way ANOVA. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05; [versus C] cc = *p* < 0.01; all data sets *n* ≥ 5. Blank box = unimpacted control (C), shaded boxes = traumatized cartilage explants (T).

#### *2.5. Trauma-Induced Expression of Aggrecanases and Subsequent Aggrecan Degradation Are Largely Decreased after Treatment with Glucosamines or PUGNAc*

≥ 5. Blank box

Influence of the HBP on trauma-induced expression and proteolytic activity of aggrecanases was determined by the gene expression analysis of ADAMTS4 and 5, estimation of the aggrecanase activity, and the histological assessment of PG by means of Safranin-O staining.

After cartilage trauma, gene expression levels of ADAMTS-4 and -5 were significantly enhanced ([vs. C] ADAMTS4: 7.4-fold; ADAMTS5: 3-fold; Figure 7A,B). This was reflected in increased aggrecanase activity ([vs. C] 2.9-fold; Figure 7C), which can be considered proportional to the total aggrecanase amount. Additionally, histological assessment of PG by means of Saf-O staining confirmed the increased loss of PG in traumatized cartilage explants (Figure 7D). Treatment with glucosamines or PUGNAc significantly suppressed trauma-induced gene expression of aggrecanases, as well as aggrecanase activity ([vs. T] GlcNAc: −4.8-fold; GS: −3.5-fold; PUGNAc: −4.9-fold). These anti-catabolic effects and the subsequent preservation of PG could be confirmed in the corresponding Saf-O staining of GlcNAc and PUGNAc treated cartilage 14 days after trauma. In contrast, AZA had no

additional effect on the expression levels of aggrecanases after trauma, though resulted in strong PG depletion, as demonstrated in the severe de-staining of the cartilage matrix.

−4 −3 −4

**Figure 7.** Effects of glucosamine treatment or addition of inhibitors on trauma-induced expression of aggrecanase – ≥ 5. Blank box **Figure 7.** Effects of glucosamine treatment or addition of inhibitors on trauma-induced expression of aggrecanases and PG breakdown 7 days and 14 days after trauma. Catabolic processes were assessed by means of gene expression analysis of (**A**) ADAMTS-4 and (**B**) ADAMTS-5, as well as (**C**) quantification of relative aggrecanase activity (7 days after trauma). Moreover, (**D**) PG content in cartilage explants 14 days after trauma and corresponding treatments were evaluated by means of exemplary images of cartilage after Saf-O staining. Statistical analysis: (**A**–**C**) one-way ANOVA. Significant differences between groups were depicted as: [versus T] \* = *p* < 0.05, \*\* = *p* < 0.01; [versus C] c = *p* < 0.05, cc = *p* < 0.01; all data sets *n* ≥ 5. Blank box = unimpacted control (C), shaded boxes = traumatized cartilage explants (T).

#### **3. Discussion**

The HBP is essentially responsible for the generation of UDP-GlcNAc, the building block for posttranscriptional O-GlcNAcylation, which is not only involved in the biosynthesis of PG but also in the regulation of various metabolic processes [9,10]. Despite its crucial role in cartilage homeostasis, the actual implication of the HBP during OA development and, in particular, after traumatic cartilage injury remains largely unknown.

In the present study, we investigated the impact of targeted manipulation of the HBP in various ways. First, we added two different glucosamine derivatives, GS and GlcNAc, in order to increase the substrate availability and potentially boost HBP activity. Second, we inhibited the enzymatic activity of OGA via PUGNAc, thus impairing reversibility of O-GlcNAcylation and causing accumulation of O-glycosylated proteins. Third, we directly inhibited the rate-limiting enzyme of the HBP, GFAT-1, by means of AZA in order to suppress the generation of UDP-GlcNAc and therefore reducing subsequent O-GlcNAcylation. Unlike previous studies, which mainly focus on glucosamine administration as a therapeutic approach against advanced and symptomatic OA, this is the first study evaluating targeted modulation of the HBP after a singular cartilage impact using a human ex vivo trauma model. The experimental design allows investigation of HBP-related processes as a possible therapeutic target after mechanical injury and subsequent cartilage degeneration on cellular and molecular level, which in vivo might promote the development of a PTOA.

Some years ago, glucosamines, and in particular GS, have been propagated to be an ideal therapeutic approach in OA treatment, combining both pain relief and cartilage regeneration. Accordingly, glucosamine application was highly recommended in the guidelines of the European League Against Rheumatism (EULAR) for the management of knee OA in 2003 [21]. In fact, different reviews and meta-analyses between 2003 and 2008 summarized the outcomes of various clinical trials on GS and concluded that it was effective in pain reduction, improvement of the joint functionality, as well as the overall reduction of OA progression and subsequent risk of joint replacement [22]. About 10 years later, the Osteoarthritis Research Society (OARSI) guidelines stated the recommendation about glucosamine treatment as "uncertain" in case of symptom relief and "not appropriate" for disease modification [4]. Taken together, therapeutic efficacy of glucosamines in OA disease remains controversial, especially because of the inconsistency between study reports deriving from industry-sponsored and independent trials, as well as a generally large heterogeneity among studies [4]. Nevertheless, as mentioned above, we investigated a novel aspect in glucosamine administration and therapeutic targeting of the HBP, respectively, with a focus on attenuation of trauma-induced pathomechanisms and thus prevention of PTOA development. In our study, GlcNAc exhibited cell protective effects after cartilage trauma; however, this could not be confirmed in the case of GS, which led to enhanced cell death in doses higher than 1.5 mM. In fact, both cell protective and as cytotoxic effects of glucosamines have been previously reported. On the one hand, glucosamine has been found to induce autophagy in chondrocytes, which is considered to facilitate cell survival and protect against cartilage degeneration [20,23–25]. On the other hand, long-term exposure and high glucosamine concentrations were found to promote mitochondrial and peroxisomal dysfunction, as well as accumulation of very long chain fatty acids (VLCFA) [20]. Therefore, glucosamines might act as inducers and suppressors of autophagic and apoptotic processes at the same time, possibly depending on the respective experimental conditions. Interestingly, cytotoxic effects have been reported for glucosamines (i.e., glucosamine hydrochloride (GlcN-HCl)), [20,26,27] but not in case of GlcNAc. In contrast to GlcNAc, glucosamine has been found to be actively internalized by chondrocytes via glucose transporters (GLUTs), thus inhibiting glucose uptake and leading to depleted intracellular ATP stores [28]. Shikhman et al. supposed that GlcNAc interacts with GLUTs, although positively affecting their affinity for glucose, the main source for energy supply and precursor of GAG synthesis in chondrocytes [28].

In line with Uitterlinden et al., who reported both anti-catabolic and anti-anabolic effects of GS and GlcN-HCl in OA cartilage [16], we found significant chondroprotective effects for GlcNAc and GS after cartilage trauma, but also observed an unexpected decline in CPII. On first sight, this finding contradicts the general opinion that glucosamine enhances the ECM synthesis in cartilage; however, post-translational modification of type II collagen does not depend upon the HBP or UDP-GlcNAc, respectively [29]. Therefore, the suppressive effects of glucosamines clearly deserve further investigation. While GS suppressed the gene expression of ACAN and HAS2, GlcNAc increased the corresponding

mRNA levels to some extent, depending on the concentration. In accordance with this, GlcNAc has previously been found to accelerate the expression of HAS2 and subsequent synthesis of hyaluronan, while glucosamine resulted in opposite effects [16,28]. The differential impact of GlcNAc and GS on chondroanabolism might also account for divergent findings concerning the PG content, as demonstrated by Saf-O staining. Despite similar suppression of aggrecanases and MMPs found for both glucosamine derivatives, higher PG decline was found in GS treated cartilage explants compared to GlcNAc treated explants.

Anti-catabolic effects of glucosamines have been previously reported, though mainly addressing interleukin 1b (IL-1b)-mediated MMP and ADAMTS expression [14,30,31]. Nevertheless, similar to cytokine-induced catabolism, the therapeutic effects of glucosamine administration after cartilage trauma might result from the inhibition of OA-associated mitogen-activated protein kinase (MAPK) pathways, including c-Jun N-terminal kinase and p38, as shown for GS [32]. However, inhibition of IL-1b-induced catabolism by GlcNAc has not been found to have any effect on ERK, JNK, or p38 MAPK pathways [33]. Moreover, spontaneous (i.e., age-dependent) and induced (i.e., via OGA-inhibitor thiamet-G) accumulation of O-GlcNAcylated proteins has even been associated with enhanced MAPK phosphorylation [34,35]. This activation correspondingly induced both MMP expression and chondrogenic differentiation of ATDC5 cells at the same time [34]. These contradictory observations imply a dual effect of O-GlcNAcylation on MAPK signaling, which clearly deserves further investigation with regard to cartilage trauma and OA development.

In our study, PUGNAc administration resulted not only in cell and chondroprotective effects but also in chondroanabolic effects after cartilage trauma, implying a positive effect by OGA-inhibition and subsequent accumulation of O-glycosylated proteins. In line with this, inhibition of GFAT-1 by means of AZA was found to result in opposite effects. Hence, we concluded that HBP functionality and O-GlcNAcylation, respectively, is essential for cartilage homeostasis and provides protection against trauma-induced pathomechanisms. Accordingly, expression of OGT and O-GlcNAcylation is thought to be responsive to cellular stress, comprising oxidative stress, ER stress, ischemia reperfusion injury, and more; however, only little is known about regulation of OGA in injured cells [36,37]. Stressinduced upregulation of OGT leads to enhanced O-GlcNAc levels and can be considered a pro-survival signaling program, while, in contrast, reduced O-GlcNAcylation increases susceptibility of cells and tissues to injury [36,38,39]. Surprisingly, cartilage trauma did not induce the gene expression of OGT but OGA, which was also observed in highly degenerated tissue (ICRS score ≥ 3). Comparable findings were described by Tardio et al., who determined a similar alteration of the OGA to OGT ratio in cartilage of OA patients, as well as after IL-1b stimulation of isolated OA chondrocytes [13]. It might be possible that the responsiveness of this pro-survival signaling program decreases with age. In fact, aging has a high impact on protein O-GlcNAcylation and vice-versa. Accumulation of O-GlcNAc modified proteins has been associated with development and progression of various age-related diseases and was found in OA cartilage tissue, despite enhanced OGA levels [12,13]. Comparable accumulation of O-GlcNAc modified proteins has also been described in aged retina; however, though, the addition of PUGNAc or UDP-GlcNAc and subsequent enhancement of O-GlcNAcylation reduced intracellular oxidative stress levels [35], which corroborates our findings.

Direct inhibition of GFAT-1 by AZA, and thus attenuation of all HBP-associated processes, resulted in enhanced expression of catabolic enzymes and subsequent ECM destruction after cartilage trauma. This was demonstrated by increased concentrations of type II collagen breakdown product C2C and severe PG depletion in Saf-O stained cartilage explants. However, loss of PG in AZA-treated cartilage explants might also be a result of both enhancement of protease activity and decrease of ECM synthesis, as shown in reduced gene expression of COL2A1, HAS2, and ACAN, as well as type II collagen synthesis. There is only little known about the impact of GFAT-1 inhibition by AZA or any other component in cartilage; however, Honda et al. reported significant impairment in the synthesis of cartilage-characteristic PG-H in chicken embryos exposed to AZA [40].

#### **4. Materials and Methods**

#### *4.1. Specimen Preparation and Cultivation Conditions*

Overall, macroscopically intact tissue samples (International Cartilage Repair Society (ICRS) score ≤ 1) from femoral condyles of 15 patients (mean age 63 years; 8 male and 7 female patients) were included in the study. Human cartilage was obtained from donors undergoing total knee joint replacement due to OA. Informed consent was obtained from all patients according to the terms of the Ethics Committee of the University of Ulm. Fullthickness cartilage explants (Ø = 6 mm) were harvested, weighed, and cultivated in serumcontaining medium (1:1 DMEM/Ham's F12 supplemented with 10% fetal bovine serum, 0.5% penicillin/streptomycin (PAA Laboratories, Pasching, Austria), 0.5% L-glutamine, and 10 µg/mL 2-phospho-L-ascorbic acid trisodium salt) for 24 h in an incubator (37 ◦C, 5% CO2, 95% humidity). Afterwards, the explants were traumatized and cultivated for 7 days and 14 days (only for histological assessment), respectively, in serum-free medium (DMEM supplemented with 1% sodium pyruvate, 0.5% L-glutamine, 1% non-essential amino acids, 0.5% penicillin/streptomycin, 10 µg/mL 2-phospho-L-ascorbic acid trisodium salt, and 0.1% insulin-transferrin-sodium selenite (Sigma-Aldrich, Taufkirchen, Germany)). All chemicals were purchased from Biochrom (Berlin, Germany), unless specified otherwise. Additionally, highly degenerated (ICRS score ≥ 3) and corresponding macroscopically intact tissue samples (ICRS grade ≤ 1) of 7 patients (mean age 67 years) were immediately snap frozen for RNA isolation to evaluate degeneration-associated changes in gene expression.

#### *4.2. Impact Loading and Subsequent Treatment*

The cartilage explants were subjected to a single impact load of 0.59 J using a droptower model, as previously described [41], and further cultivated under serum-free conditions (described above). Unloaded explants served as controls. Impacted cartilage explants were treated with the following additives (Sigma-Aldrich) for 7 and 14 days, respectively: GlcNAc or GS (each 0.1–10 mM), an inhibitor of O-GlcNAcase (PUGNAc (O- (2-acetamido-2deoxy-D-glucopyranosylidene)amino-N-phenylcarbamate): 0.05–0.15 mM) or the GFAT1-inhibitor azaserine (AZA: 0.005–0.1 mM) (Figure 1). Fresh additives were added concomitantly with medium change every 2–3 days.

#### *4.3. Live/Dead Cell Cytotoxicity Assay*

To determine the percentage of viable cells 7 days after trauma, a live/dead viability/cytotoxicity assay (Molecular Probes, Invitrogen) was performed, as previously described [42]. In short, unfixed tissue sections (0.5 mm thickness) were stained with 1 µM calcein AM and 2 µM ethidium homodimer-1 for 30 min. After washing in PBS, they were microscopically analyzed by means of a z-stack module (software AxioVision, Carl Zeiss, Jena, Germany). Three pictures were made from each tissue section. All cells on the picture were counted manually (Image J software version 1.42q). The average count per picture was about 2000 cells.

#### *4.4. mRNA Isolation and cDNA Synthesis*

For total RNA isolation 7 days after trauma, cryopreserved cartilage explants were pulverized with a microdismembrator S (B. Braun Biotech, Melsungen, Germany). Subsequently, RNA was isolated using the Lipid Tissue Mini Kit (Qiagen, Hilden, Germany). RNA was reverse transcribed with the Omniscript RT Kit (Qiagen) and used for quantitative real-time PCR analysis (StepOne-PlusTM Real-Time PCR System, Applied Biosystems, Darmstadt, Germany).

#### *4.5. Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)*

Determination of the relative expression levels was performed by means of qRT-PCR analysis using the ∆∆ Ct method. To detect desired sequences, a TaqManTM Gene Expression Master Mix for TaqMan Gene Expression Assay (both Applied Biosystems) was used for the fol-

lowing probes: Hs00192708\_m1 (ADAMTS4), Hs00199841\_m1 (ADAMTS5), Hs00264051\_m1 (COL2A1), Hs00899865\_m1 (GFAT-1), Hs00193435\_m1 (HAS2), Hs02800695\_m1 (HPRT1), Hs00899658 (MMP-1), Hs00233992\_m1 (MMP-13), Hs01028844\_m1 (OGA/MGEA5), and Hs00269228\_m1 (OGT). Power SYBR Green PCR Master Mix (Applied Biosystems) was used for 18S rRNA, 5′ - CGCAGCTAGGAATAATGGAATAGG-3′ (forward), 5′ -CATGGCCT CAGTT CCGAAA-3′ (reverse), and Platinum SYBR Green qPCR SuperMix-UDG (Invitrogen, Darmstadt, Germany) for GAPDH, 5′ -TGGTATCGTGGAAGGAC TCATG-3′ (forward), and 5′ -TCTTCTGGGTGGCAGTGATG-3′ (reverse). mRNA expression was determined by normalizing the expression levels separately to the endogenous controls (18S rRNA, GAPDH, and HPRT1) and subsequently calculating the ratio mean values in relation to the gene expression level of the untreated, unimpacted control.

#### *4.6. Culture Media Analysis by Means of Commercial ELISA Kits*

Biomarker release into culture media (7 days after trauma) was evaluated by means of enzyme-linked immunosorbent assays (ELISAs): secreted MMP-13 was determined using the Human Quantikine ELISA kit (Ray-Biotech, Norcross, GA, USA). Evaluation of type II collagen synthesis was performed using a CPII ELISA (Ibex, Québec, QC, Canada). The assay quantified type II collagen carboxy propeptide (CP II) cleaved from pro-collagen II after its release into the matrix and directly correlated with newly synthetized type II collagen. Degradation of type II collagen was measured using a C2C ELISA (Ibex), detecting a neoepitope generated during collagenase-mediated breakdown of type II collagen. Aggrecanase activity was measured using the Sensitive Aggrecanase Activity Assay from Biotez (Berlin, Germany) for serum-free cell culture supernatants. The kit consists of two modules. First, the substrate (modified aggrecan interglobular domain) is proteolytically cleaved by sample-derived aggrecanases, releasing a peptide with a N-terminal ARGSVIL sequence. This peptide can then be quantified via the specific ELISA module. In principle, the assay quantifies the total aggrecanase activity, comprising that of ADAMTS-1, -4, and -5.

The total amounts of MMP-13, C2C, and CP II, as well as the aggrecanase activity, were relativized on the weight multiplied by cell viability of the corresponding cartilage explant [41,42].

#### *4.7. Gelatin Zymography*

Quantification of pro-MMP-2 and active MMP-2 was performed by gelatin zymography, as previously described [41]. In short, culture media (7 days after trauma) were mixed 1:2 with nonreducing zymogram sample buffer (Bio-Rad, Munich, Germany), loaded onto 10% polyacrylamide gels (Carl Roth, Karlsruhe, Germany) containing 2 mg/mL gelatin (Merck, Darmstadt, Deutschland). After electrophoresis (Mini-PROTEAN Tetra Cell System, Bio-Rad), the gels were washed in zymogram renaturation buffer twice for 15 min and incubated in zymogram development buffer for 20 h at 37 ◦C (both Bio-Rad). Staining with Coomassie solution and subsequent destaining revealed clear bands originating from MMP activity. Band intensities (INT\*mm<sup>2</sup> ) were quantified with the Geldoc XR system (Bio-Rad) and relativized on tissue weight and cell viability, as mentioned above. An internal standard (positive control) was run on each gel and used to reduce the inter assay variance.

#### *4.8. Safranin-O Staining*

To evaluate the content of PG within the cartilage tissue, appropriate histological analysis was performed exemplarily (*n* = 1) 14 days after trauma. In short, explants were fixed (4% paraformaldehyde) and embedded in paraffin. Dewaxed and rehydrated sections (3.5 µm) were stained with SafO (Thermo Fisher Scientific, Schwerte, Germany) and Fast Green (Sigma-Aldrich), followed by a final staining of the cell nuclei by Gill's hematoxylin No. 3 (Sigma-Aldrich) and documentation with an Axioskop 2 mot plus (Zeiss, Oberkochen, Germany).

#### *4.9. Statistical Analysis*

Experiments were analyzed using GraphPad Prism8 (GraphPad Software Inc., La Jolla, CA, USA). Each data point represented an individual donor (biological replicate); technical replicates from the same donor were not performed. Data sets with *n* ≥ 5 were tested for outliers by means of the Grubbs outlier test. Outliers were not included in statistical analyses. For parametric data sets, a one-way analysis of variance (ANOVA) with the Sidak post-test was used. Nonparametric data sets were analyzed by means of a Kruskal– Wallis test with Dunn's post-test. For data sets derived from highly degenerated tissue (ICRS grade ≥ 3), an unpaired two-tailed t-test was performed. The significant level was set to a = 0.05. Values in diagrams are given as boxplots (median; whiskers: min to max).

#### **5. Conclusions**

In conclusion, our results demonstrate the overall importance of the HBP in cartilage homeostasis and degeneration processes, which could be significantly influenced by targeted modification. In this context, we observed predominately chondroprotective and chondroanabolic effects associated with O-GlcNAcylation and the HBP after ex vivo cartilage trauma, implying that alteration of the glycosylation profile might have a detrimental impact on cartilage homeostasis. However, it cannot be excluded that excessive accumulation of O-GlcNAc modified proteins in age might negatively affect cartilage homeostasis in the same manner as its decline, as confirmed by the alteration in the gene expression of HBP-related enzymes in ex vivo traumatized macroscopically intact and in vivo highly degenerated OA cartilage. So far, it is not known whether this reflects a primary pathogenic event or an insufficient protective mechanism in aging and OA, respectively. With respect to various studies reporting contradictory effects of enhanced O-GlcNAcylation in aging, we conclude that this post-transcriptional modification might be a two-edged sword, in particular regarding degenerative diseases. However, in the early post-traumatic situation, the HBP may represent a promising therapeutic target.

**Author Contributions:** Conceptualization, R.E.B. and J.R.; validation, J.R. and J.B.; investigation, J.R. and J.B.; writing—original draft preparation, J.R.; writing—review and editing, R.E.B., J.B., J.R. and F.Z.; visualization, J.R.; project administration, R.E.B. and J.R.; funding acquisition, R.E.B., J.R. and F.Z. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research study was funded by the Deutsche Initiative Arthroseforschung of the German Society of Orthopedics and Orthopedic Surgery (DGOOC). Moreover, this project was supported by the European Social Fund and by the Ministry of Science, Research and Arts Baden-Württemberg.

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki and approved by the Ethical Committee of the University of Ulm, Germany (ethical approval number 353/18).

**Informed Consent Statement:** Informed consent was obtained from all subjects involved in the study.

**Data Availability Statement:** Data are contained within the article.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**

## *Article* **A Novel Method Facilitating the Simple and Low-Cost Preparation of Human Osteochondral Slice Explants for Large-Scale Native Tissue Analysis**

**Jacob Spinnen 1, \* , Lennard K. Shopperly 1 , Carsten Rendenbach 2 , Anja A. Kühl 3 , Ufuk Sentürk 4 , Daniel Kendoff 5 , Shabnam Hemmati-Sadeghi 1 , Michael Sittinger <sup>1</sup> and Tilo Dehne 1**

**Abstract:** For in vitro modeling of human joints, osteochondral explants represent an acceptable compromise between conventional cell culture and animal models. However, the scarcity of native human joint tissue poses a challenge for experiments requiring high numbers of samples and makes the method rather unsuitable for toxicity analyses and dosing studies. To scale their application, we developed a novel method that allows the preparation of up to 100 explant cultures from a single human sample with a simple setup. Explants were cultured for 21 days, stimulated with TNF-α or TGF-β3, and analyzed for cell viability, gene expression and histological changes. Tissue cell viability remained stable at >90% for three weeks. Proteoglycan levels and gene expression of *COL2A1*, *ACAN* and *COMP* were maintained for 14 days before decreasing. TNF-α and TGF-β3 caused dosedependent changes in cartilage marker gene expression as early as 7 days. Histologically, cultures under TNF-α stimulation showed a 32% reduction in proteoglycans, detachment of collagen fibers and cell swelling after 7 days. In conclusion, thin osteochondral slice cultures behaved analogously to conventional punch explants despite cell stress exerted during fabrication. In pharmacological testing, both the shorter diffusion distance and the lack of need for serum in the culture suggest a positive effect on sensitivity. The ease of fabrication and the scalability of the sample number make this manufacturing method a promising platform for large-scale preclinical testing in joint research.

**Keywords:** osteoarthritis; osteochondral explant culture; joint modelling; pharmacological assay; native tissue analysis

#### **1. Introduction**

Osteoarthritis (OA) is a condition involving the degeneration of articular cartilage, sclerosis of subchondral bone and chronic inflammation of the synovial membrane. To date, it is the main cause of physical disability worldwide [1,2]. Both the underlying pathomechanisms of OA as well as the mechanisms of physiological reorganization of cartilage tissue are poorly understood. Due to this lack of understanding, the treatment of such defects

**Citation:** Spinnen, J.; Shopperly, L.K.; Rendenbach, C.; Kühl, A.A.; Sentürk, U.; Kendoff, D.; Hemmati-Sadeghi, S.; Sittinger, M.; Dehne, T. A Novel Method Facilitating the Simple and Low-Cost Preparation of Human Osteochondral Slice Explants for Large-Scale Native Tissue Analysis. *Int. J. Mol. Sci.* **2021**, *22*, 6394. https://doi.org/10.3390/ijms22126394

Academic Editor: Lih Kuo

Received: 25 May 2021 Accepted: 9 June 2021 Published: 15 June 2021

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remains challenging. However, research in recent years has provided increasing evidence that cartilage cells have the general ability to regenerate if stimulated correctly. Limited traumatic cartilage defects can now be successfully treated and regenerated by autologous chondrocyte transplantation and specific cell-modulating substances. The emergence of growth factors as therapeutic agents is expected to further enable the regeneration of osteochondral tissue [3–5], increasing the need for an adequate platform to assess cartilage (repair) treatment strategies.

Traditionally, two-dimensional chondrocyte cultures in monolayer and OA animal models have been the main tools available for preclinical testing of the efficacy and side effects (ADME (absorption, distribution, metabolism, excretion) screening) of such substances. Unfortunately, the former often leads to limited insights due to the lack of representation of the complex tissue composition of the joint, while the latter, in addition to ethical aspects, entails high costs and time expenditure and is therefore usually only applied when a basic efficacy already seems very likely [6,7]. Recently, this spectrum has been expanded to include sophisticated 3D cultures and organ-on-chip applications. However, 3D cultures are either effortful to maintain or, in the case of joint-on-a-chip, not yet commercially available. Therefore, rapid, agile research and development of regenerative therapeutics for cartilage regeneration is limited to a certain extent [8–10].

A sustainable middle ground in joint modelling is the use of tissue explants. Explants are living, native and functional parts from organs, which are obtained from donor tissues or cadavers. Explants can accurately represent part of the tissue composition and architecture. Specifically, the tissue cells remain in their native extracellular matrix (ECM) configuration and questions regarding their response to biological stimuli can be answered much more reliably in the native situation than, for example, with monolayer cultures of only one cell type [11–13]. However, this modelling technique is severely limited by the availability of donor tissue. Employing the frequently used technique of vertical punching of the tissue and subsequential production of osteochondral punchings with diameters of at least 3 mm, only a very small number of explants can be produced from a tissue sample, depending on the tissue properties. Furthermore, the diffusion of nutrients and potentially interacting proteins is severely impeded by the amount of glycosaminoglycans in the tissue [14]. Hence, the explant thickness negatively correlates with information that can be obtained about the behavior of cells located deeper in the tissue.

To address this issue, we have developed a low-cost and uncomplex method to produce explant cultures using a thin-section approach. Similar to the live-slice technology known from neurophysiology, the method allows the production of vital tissue slices with a thickness of 500–800 µm [15]. This allows the production of up to 100 slice cultures from a single tissue sample for subsequential analysis of many different substances and concentrations. Furthermore, the thin tissue architecture is suitable for easy-to-maintain semi-static culture conditions, as the perfusion distance is sufficiently short to reliably guide both nutrients and any therapeutic agents through the tissue to the target cells.

This study aimed to analyze the suitability of osteochondral slices for osteochondral tissue modelling and pharmacological screening of biologically active substances. For this purpose, we obtained a large number of human slice cultures from surgically explanted tibial plateaus and analyzed the behavior of the tissue over 21 days at the histological, metabolic and transcriptional levels. Furthermore, we examined the reactivity of the embedded chondrocytes to the factors TNF-α and TGF-β3, known to affect cartilage matrix synthesis, to determine whether the tissue is in a sufficiently close state to that of the native situation to be used as a testing device [16,17].

#### **2. Results**

To develop a physiological model for osteochondral tissue which allows for quick handling and high-throughput applications with limited donor material at hand, we developed a method for the manufacturing of native osteochondral live slice cultures from human joint tissue. 500–800 µm thick osteochondral slice cultures were prepared

from 23 different surgically explanted tibial plateaus using a custom-made microtome insert and then cultured in hanging inserts of 24-well plates. In this study, we assessed whether the explanted live slice cultures maintain their physiological properties such as long-term cell viability, gene expression, extracellular matrix and responsiveness towards biological stimuli.

#### *2.1. Suitability for Slicing Varies with the Degree of Subchondral Bone Sclerosis*

23 of 25 donors were suitable for the production of slice explants. The extent of subchondral sclerosis of the bone proved to be a decisive factor for the suitability of slice culture production. The brittleness of the bone increases sharply with the degree of sclerosis, leading to the subchondral cancellous bone not being cut smoothly by the impact of the blade but instead being crushed. Three punch cylinders were easily obtained from all other donors, from each of which 30–36 slice cultures could be safely produced. For reasons of logistical feasibility, we only cultivated the slice cultures from one punch at a time-from a technical perspective, the cultivation of 90–100 slice cultures per donor would have been possible.

#### *2.2. Confocal Laser Scanning Microscopy Shows Highly Conserved Spatial Cell Order*

To validate the use of the resazurin assay and to evaluate the spatial distribution of living and dead cells, additional live/dead determination via CLSM was performed on 13 slices of two donors (7 and 8) after three weeks of culture. The slice explant cultures were thin enough to penetrate deep into the tissue layers with confocal laser microscopy and produce a three-dimensional image of the cell distribution- and vitality (3D rendered video in supplemental files). The analysis revealed >90% viability in non-heated slices, roughly 50% viability in slices that were heated for 30 min at 60 ◦C, and <10% chondrocyte viability after 60 min of heating (Figure 1a). Chondrocytes remained localized in their cartilage-typical, spatial alignment (column-like deep zone; pearl-bead structure in the mid-zone; tightly packed cells in the superficial zone, Figure 1a). No cell culture effects due to nutrition gradients were observed (e.g., no elongated cells in the peripheral tissue).

#### *2.3. Resazurin Assay Correlates with Optically Determined Viability*

To ensure that the resazurin assay sufficiently reflects viability and is not obstructed by the dense osteochondral ECM, samples of varying viability (obtained by heating at 60 ◦C) were analyzed with the resazurin metabolic assay and by live/dead determination via PI/FDA staining and CLSM analysis. The results of both methods were correlated, as shown by the correlation coefficient of R<sup>2</sup> = 80–84.5% (Figure 1b). We regarded this as sufficient to assess the general viability state of the tissue culture. Further viability analyses of tissue cultures were therefore performed using the resazurin assay since it allows for progredient viability analysis of the same tissue slice in a non-destructible manner.

#### *2.4. Cell Viability Is Maintained in F*<sup>−</sup> *and Increased in F<sup>+</sup> Groups over 21 Days*

After 7 days, F<sup>−</sup> cultured slices of the cytokine-stimulated groups exhibited either a slight increase or maintenance of viability as determined by the resazurin assay. Stimulation with TNF-α resulted in 101 ± 23% (10 ng/mL) and 106 ± 31% (40 ng/mL) compared to control, while TGF-β3 stimulated slices exhibited an increase to 113 ± 26%. Unstimulated slices showed a slight decrease to 91 ± 34%. No significant differences in viability from day 0 to day 7 as well as between the four different cytokine stimulation groups were observed. Addition of FBS to the culture medium resulted in significantly increased viability values in all stimulated groups (TNF-α: 214 ± 58% (low), 227 ± 84% (high); TGF-β3: 218 ± 48%) and the control (231 ± 50%) (Figure 1c(i)). During three-week culture without FBS viability values of 114 ± 43, 108 ± 52% and 113 ± 55% were observed after 7, 14 and 21 days, respectively. Statistical analysis revealed no significant differences in viability between the timepoints (Figure 1c(ii)).

α or with 20 ng/mL TGF β3, **Figure 1.** Viability analysis of osteochondral live slices. (**a**) Confocal laser scanning microscopy of slices at three weeks of culture following heating at 60 ◦C for 0, 30 or 60 min and live/dead staining with PI/FDA. Vital cells appear green, while dead cells appear red. (**b**) Correlation analysis of vital cell count as determined by CLSM and metabolic activity as determined by AlamarBlue resazurin assay. Two donors and a total of 13 slices with varying viability were analyzed and revealed correlation coefficients of 0.8 and 0.845. (**c**) Resazurin assay results with and without additional stimulation. i: Viability of slices on day 7 relative to day 0. Slices were stimulated with 10 or 40 ng/mL TNF-α or with 20 ng/mL TGF-β3, and either with or without FBS. ii: Viability of slices without FBS or other stimulating factors over 21 days, relative to day 0.

#### *2.5. Expression of Cartilage-Typical Markers ACAN and COL2A1 Is Maintained for 14 Days in Unstimulated Slice Culture*

– Expression of cartilage-relevant genes *COL2A1* and *ACAN* remained stable relative to their day 0 value for 14 days and declined significantly by day 21 in unstimulated slice cultures (no stimulating factors, no FBS) (*COL2A1*: 32 ± 23%, *p* < 0.001; *ACAN:* 51 ± 64%, *p* < 0.05; Figure 2a). Cartilage remodeling marker *COL1A1* showed a short increase after 7 days but returned to levels comparable to day 0 by day 21. Expression of *COMP* decreased sooner and significantly to 45 ± 30% after 7 days and further to 27 ± 21% and 14 ± 12% after 14 and 21 days (all *p*-values < 0.001).

β3 stimulated slices exhibited an increase to 113 ± 26%. Unstimulated

α: 214 ± 58% (low), 227 ± 84% (high); TGF β3:

α resulted in 101 ± 23% (10 ng

−

*−*

− α or TGF β3 and with or − α or TGF β **Figure 2.** Gene expression analysis of live slice explants via RT-qPCR. (**a**) Gene expression of cartilage-typical markers *COL2A1*, *COL1A1*, *ACAN* and *COMP* in F <sup>−</sup> cultured slices over three weeks (*n* = 12) relative to the day 0 value. (**b**) Gene expression of the same cartilage-typical markers as in A, after 7 days of stimulation with TNF-α or TGF-β3 and with or without FBS (*n* = 6) relative to the control on day 7. (**c**) Expression of proliferation marker *Ki-67* in F <sup>−</sup> cultured slices over three weeks (*n* = 12) relative to the day 0 value and (**d**) after 7 days of stimulation with TNF-α or TGF-β3 and with or without FBS (*n* = 6), relative to the day 0 control to show the larger increase in the F <sup>+</sup> group. Statistically significant differences denoted as \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

#### *− 2.6. Transcription of ECM Proteins Remains Highly Reactive to External Stimuli in F* <sup>−</sup> *Culture*

α β3 to either − − α To test cell reactivity to external stimuli, we intended to induce measurable changes in chondrocyte gene expression. Slices were stimulated with TNF-α and TGF-β3 to either suppress or induce the expression of cartilage-relevant gene markers. Gene expression analysis showed differing results in the F <sup>−</sup> and F + stimulation groups. In the F <sup>−</sup> group, stimulation with TNF-α resulted in significant and dose-dependent reductions of *COL2A1, ACAN* and *COMP* expression. Compared to the day 7 control, *COL2A1* expression exhibited levels of 11 ± 7% and 5 ± 5% in low and high dose TNF-α stimulations, respectively. *ACAN* expression also was significantly lower at 42 ± 33% and 15 ± 12%, while *COMP* expression dropped to 44 ± 48% and 10 ± 14%. Stimulation with TGF-β3 resulted in a 19 ± 25-fold higher *COL1A1* expression and 9 ± 5-fold higher *COMP* expression (Figure 2b). Expression of *MMP13* did not change significantly after incubation with TNF-α but was significantly lower after stimulation with TGF-β3 (6 ± 6%, Supplemental Figure S1). In the F <sup>+</sup> group, stimulation with TNF-α resulted in a reduced *COL2A1* expression of 33 ± 11% in the lower concentration and 11 ± 4% in the high concentration compared to the control. Mean expression levels of *ACAN* and *COMP* in F <sup>+</sup> also decreased after TNF-α exposure, although no coherent dose-dependent or significant effects could be observed. TGF-β3 stimulation resulted in a 29 ± 25-fold higher expression of *COMP* and a 2.4 ± 2.7-fold

higher *COL1A1* expression (Figure 2b), as well as a decrease to 42 ± 40% of *MMP13* expression (Supplemental Figure S1).

#### *2.7. Addition of FBS to Culture Media Causes Dominant Effects on Proliferation Marker Ki-67 Expression*

Expression of *Ki-67* as a marker for cell proliferation in serum-free culture varied strongly over three weeks and between donors, as shown by the very high standard deviations and surge to a >1000-fold increase on day 14 before it returned to a 28 ± 90-fold higher level compared to day 0 on day 21 (Figure 2c). Even though the average increase in *Ki-67* expression from day 0 to 7 was substantial, presumably because of the low absolute initial expression of *Ki-67* at day 0, very high variability between donors was observed and led to statistically insignificant results. Stimulation with low concentration TNF-α in F <sup>−</sup> groups led to 31 ± 65% of day 0 expression, while unexpectedly higher concentrated TNF-α resulted in a 50 ± 112-fold increase. TGF-β3 stimulation revealed a 37 ± 60-fold increase in expression after 7 days compared to day 0 control (*p* = 0.06). In the F<sup>+</sup> group, however, all 3 stimulation groups and the control exhibited statistically significant changes between day 0 and 7, reaching fold-changes compared to day 0 of 12,045 ± 17,483 (TNF-α low), 1686 ± 2180 (TNF-α high) and 4889 ± 9110 (TGF-β3). No statistically significant differences were detected among the F<sup>+</sup> cultured stimulation groups or between either of the groups and the day 7 negative control (Figure 2d).

#### *2.8. Tissue Slices React to TNF-α Stimulation with Observable Remodeling of ECM and Cellular Swelling*

To analyze whether native slice cultures retain their tissue-specific histological composition or whether they can respond to molecular stimuli with remodeling of the ECM, the slices of long-term culture and those of TNF-α-stimulated cultures were stained with Safranin-O, because TNF-α is an important inducer of matrix degeneration processes in cartilage. The proteoglycan contents of non-stimulated, F<sup>−</sup> cultured slices on days 0, 7, 14 and 21 (Figure 3a), as well as TNF-α-stimulated slices and negative controls on days 0 and 7 (Figure 3b), were quantified histomorphometrically. Even though a statistically significant decrease in proteoglycan content (*p* < 0.05) compared to day 0 was observed on day 7 in the long-term setup (Figure 3a(ii)), the mean proteoglycan content overall was maintained, as on days 14 and 21 it did not significantly differ from day 0. Proteoglycan content in the TNF-α-treated group was significantly lower than in the untreated control on day 7 (68 ± 24%; *p* < 0.05; Figure 3b(ii)). Corresponding to the changes in gene expression, TNF-α-stimulated slices exhibited a strongly altered histomorphology. Intra-cartilage fibers appeared less dense, while chondrocyte diameter and apparent volume increased (Figure 3c).

α α α **Figure 3.** Histomorphometrical and histological analysis of live slices. (**a**) Safranin-O-stained slices of one donor (i) and histomorphometrical analysis (ii) over three weeks. (**b**) Comparison of staining intensity in TNF-α-stimulated and nonstimulated slices. Stimulated slices (three exemplary donors shown in (i)) show a mean reduction in red intensity of 32% (ii, *n* = 6). (**c**) Representative picture of TNF-α-treated slice and control. TNF-α-treated slices show less intense Safranin-O staining, loosened matrix structure and swollen cells in the mid-zone of the cartilage. Statistically significant differences denoted as \* *p* < 0.05.

#### **3. Discussion**

The herein presented cost- and time-effective setup of surgical punch, 3D-printed microtome insert and rotary microtome is a reliable method to produce up to 100 explant cultures from a single human tissue sample. Critical to the usability of these thin osteochondral slice cultures is the long-term preservation of native tissue configuration and cell vitality despite thermal and mechanical stress development during preparation on the microtome and exposure of cells to atmospheric oxygen levels. CLSM analysis showed high cell vitality in both superficial and deeper tissue layers. Therefore, the slice thickness of 500 µm appears to be sufficient for cell protection within the tissue while ensuring adequate nutrient perfusion through motion-assisted diffusion. Furthermore, CLSM shows preservation of cartilage-specific cell architecture within the tissue even after three weeks of culture. Cartilage is divided into three distinct cellular zones: The superficial zone (densely packed with spindle-shaped cells), the middle zone (pearl cord-like alignment) and the deep zone (columnar chondrocyte alignment). Maintenance of the cellular zones depended on nutrient supply and physiological cartilage infrastructure, suggesting minimal cellular stress from malnutrition and/or hypoxia. However, as Secretan et al., already mentioned in their explant model, the disadvantage of the standard optical live/dead analysis of explants is the exclusive possibility of endpoint analysis and the uncertainty about apoptotic cells that may have been cleared in the meantime [18]. Therefore, the high correlation between cell vitality as determined by CLSM and viability determined by resazurin assay enable the utilization of the inexpensive and easy-to-handle resazurin assay for multi-point analyses in this culture form. This is particularly advantageous for high sample quantities, which can be created by this cutting method. Long-term analysis of cell viability revealed very stable viability values over 21 days with only minor fluctuations from the baseline. Interestingly, this stability was achieved in a regular cell culture medium without serum

supplementation. Most explant cultures published to date use serum mixtures between 2–10% during cultivation [18–20]. While the addition of serum is a standard cell culture procedure, it is highly desirable, especially in light of pharmacological testing, if serum were not required as an undefined and potentially interacting, interfering or confounding factor for culture maintenance. This incentive is evident from the immense increase in absolute values and variation in viability values in the short-term culture with FBS addition, where toxicity effects would presumably not be visible due to the increased metabolic activity.

In terms of long-term stability, the slice explants provide similar characteristics to conventional punch explants. Bian et al. were able to demonstrate relative stability regarding the glycosaminoglycan content in serum-free cultivation of 3 mm thick explants over several weeks [21]. While we observed a significant decrease in the histomorphometrically determined Safranin-O staining intensity on day 7, it stabilized again on days 14 and 21. This could be due to an initial outflow of glycosaminoglycans at the cut edges of the cartilage, which account for a larger proportion of the total cartilage volume than in conventional punch explants. Regarding cartilage gene expression, the results are also similar to previously published analyses of explant gene expression. Different studies also described an initial increase of aggrecan followed by a decrease after 14 days and a decrease of the collagen type II expression of >50% after three weeks of cultivation following an initial stable phase [19,22]. *COMP* as a very sensitive marker for cartilage synthesis is the only gene in the analyzed panel to show a constant, almost linear decrease over time [23,24].

Stimulation of osteochondral slices with biological stimuli showed that chondrocytes in slice cultures responded adequately to external stimulation with catabolic molecules. TNF-α is known to be a proinflammatory cytokine in the joint and to induce cartilage matrix degradation both in vivo and in vitro [25,26], but other explant models either took significantly longer for a similarly strong reduction in matrix expression and degradation of proteoglycans or required additional catabolic stimuli such as oncostatin-M or interleukin-1β [11,27,28]. Here, the short perfusion distance between the medium and cells could be an advantage for pharmacological testing, since a shorter diffusion distance results in a higher and faster penetration by the corresponding factors than in thicker explant cultures. In addition to the decrease in anabolic biomarkers such as *COL2A1*, catabolic biomarkers such as *MMP13* also responded to external stimuli. Stimulation of the cultures with TGF-β3 resulted in close to complete suppression of *MMP13* expression more clearly observed in the serum free culture. Meanwhile, stimulation with TNF-α did not significantly increase *MMP13* expression. This suggests that due to the osteoarthritic damage and semi-static nature of the culture, only decreases catabolic marker genes are suitable as indicators of efficacy in pharmacological studies.

In studies aimed at analyzing pathological processes in cartilage or the long-term effects of an external stimulus rather than large-scale screening, it would also be beneficial to examine the distribution of the different collagen subtypes histologically, as these provide the most accurate information on cartilaginous remodeling processes.

Analogous to the viability measurement, it was also shown that when serum was added to the culture, the influences of the stimuli on the chondrocytes were significantly lower. In the serum-containing group, only the decrease of *COL2A1* expression following higher-dose TNF-α stimulation and the TGF-β3-dependent increase of *COMP* on day seven reached statistical significance compared to the control. Furthermore, analysis of the proliferation marker *Ki-67* showed that in the serum-containing group, all stimulated groups including the negative control showed a huge increase, whereas, in the F<sup>−</sup> group, only TGF-β3 stimulation resulted in higher expression of *Ki-67*. This is further evidence of nonspecific responses by serum addition masking the specific effects of other stimuli, rendering it neither necessary nor advisable for pharmacological testing.

The method is limited by donor variability, the increased effort in handling, the use of priorly diseased tissue and its static nature. In comparison to conventional explants, which are merely punched out of the tissue specimen, slice cultures require several processing steps until successfully produced. Since the chondrocyte phenotype is strongly related to

mechanical stimulation, several approaches are currently aiming to increase the dynamics and longevity of explant cultures by applying mechanical pressure [13,29–31]. This approach is more suitable for thicker, cylindrical explants and would, for the thin sections, be very difficult to implement. Therefore, the native tissue statues is inherently limited to a window of 3 to 4 weeks using this preparation method. This also limits the model in particular to the analysis of toxic-catabolic processes. Anabolic stimuli such as TGF-β3, while showing an increase in matrix protein expression at the transcriptional level, are associated with the simultaneous presence of mechanical stimuli [32]. A prior induction of catabolism would most likely be required to study the effects of an anabolic stimulus as previously described by Schlichting et al. [8,33]. Furthermore, the exact reproduction of slice thickness can be impaired by different qualities of bone density. Stronger bone calcification can make it necessary to cut the slice 100–200 µm thicker as the brittleness causes the bone to crush while being cut, resulting in unusable slices. Due to these slight variations in the cutting process, every donor must be related to itself when performing multipoint analysis. Therefore, the availability of a progredient analysis method like the resazurin assay is necessary to monitor individual slice viability. In accordance with this, a proteoglycan assay from the culture supernatants would also be conceivable for the analysis of the ECM. This could also provide continuous data from a slice culture instead of having to sacrifice the culture for histological endpoint analysis.

#### **4. Materials and Methods**

#### *4.1. Sample Acquisition and Slice Production*

Whole tibial plateaus (TPs; *n* = 23; 13 female, 10 male, 61–89 years, ∅ 72 years) were acquired from patients undergoing knee arthroplasty comprising TP removal. The specimens were transported in serum-free cell culture medium (Dulbecco's Modified Eagle Medium with 1 g/L glucose; 1% Penicillin/Streptomycin; 2% HEPES, Merck, Darmstadt Germany) to the laboratory with a maximum delay of 1 h. TPs were then immediately placed in Petri dishes filled with medium prewarmed to 37 ◦C under a laminar flow cabinet. After macroscopical inspection for an osteochondral site with a well-preserved cartilage-to-bone ratio (criteria: at least 1 mm high cartilage layer; no sclerosed subchondral bone below the cartilage), an orthopedic tissue punch (OATS®, Arthrex, Naples, FL, USA) press was used to create an osteochondral cylinder punch of 20 × 10 mm (Figure 4a). The cylinder was inserted into a custom 3D-printed microtome insert (see Supplemental Figure S2 for exact insert dimensions) and then cut into eight disc-shaped cuts with a thickness of 500–800 µm using a standard rotary microtome (Cut4060, MicroTec, Brixen, Italy). An 8 mm N35 microtome blade (FEATHER, Osaka, Japan) was used for cutting, which was exchanged after the preparation of 3 osteochondral cylinders. To produce a precise cut, the rotary handle was pulled downward at the point of maximum height in a powerful swing to exert maximum force on the cutting surface and thus prevent the bone from breaking. The cuts were then split into 3 cuboid-shaped slices using a scalpel, resulting in 24–36 individual slices per punch (Figure 4b). The remains of the cylinders were then discarded.

Two slices were immediately conserved either in 5% formaldehyde or in RNAlater® (Thermo Fisher, Waltham, MA, USA), posing as day 0 samples for histological analysis and real-time quantitative polymerase chain reaction (RT-qPCR). Slices in RNAlater were subsequently stored at −80 ◦C. The remaining 22 slices were placed into hanging inserts (in a standing position with the cartilage facing up and leaning onto the upper rim of the inserts) of 24-well tissue culture plates with an 8 µm pore diameter (Transwell®, Corning, New York, NY, USA) for optimal perfusion. All treatments of the tissue were performed with sterile surgical gloves and instruments. Tissue culture plates were cultured at 37 ◦C and 5% CO<sup>2</sup> on a horizontal shaker at 10 rpm (Figure 4c).

– **Figure 4.** Fabrication and culture of osteochondral live slices. (**a**) Punching of Osteochondral cylinders. A 2 × 1 × 1 cm long osteochondral cylinder is punched out of the macroscopically unaffected area of a recently explanted human tibial plateau under sterile conditions using an orthopedic OATS ® Tissue Punch (i,ii). The resulting cylinder is shown schematically in iii, cartilaginous areas are highlighted in yellow. Front and side views of the cylinder are shown in iv. (**b**) Cutting of osteochondral slices from cylinder. The cylinder is inserted into a 3D-printed microtome insert (iv) with no additional fixation (i), and 500–800 µm thick cuts are cut out from the cylinder (ii). Fixation of the cylinder is shown schematically in iii. The resulting disc-shaped cuts (schematic: v) are cut into three parallelepipedal slices using a scalpel (vi). (**c**) Slices are immediately transferred to a hanging insert of a multi-well plate and covered in cell culture medium (i,ii). Plates are then placed on a horizontal shaker for culture at 37 ◦C and 5% CO<sup>2</sup> .

#### – *4.2. Maintenance and Stimulation*

α, 1.2 nM (40 ng α or 0.8 nM (20 ng β3 (all Peprotech, Rocky Hill, NJ Slices of donors 1–6 were divided into a serum-free (F <sup>−</sup>) culture group and a serumcontaining group (F + ). The F <sup>+</sup> group was cultured in the same medium as mentioned above with an additional 10% fetal bovine serum (FBS; Gibco, Thermo Fisher, Waltham, MA, USA). In both groups, four slices each were stimulated with either 0.3 nM (10 ng/mL) TNF-α, 1.2 nM (40 ng/mL) TNF-α or 0.8 nM (20 ng/mL) TGF-β3 (all Peprotech, Rocky Hill, NJ, USA) or no additives. Stimulating factors were pipetted with low-binding tips (Corning, New York, NY, USA) for minimal protein loss. Medium and stimulating factors were exchanged completely every three days. Analyses of slices for viability, gene expression and proteoglycan content were performed on days 0 and 7. Slices of the additional 17 donors were cultured without additional stimulating factors or FBS for 21 days in a large group viability and gene expression analysis (Figure 5). The medium was completely exchanged twice a week.

−

– – **Figure 5.** Flowchart depicting donor sample acquisition and distribution. Samples were obtained and then distributed for use in short-term (7 days, donors 1–6) stimulation experiments or long-term (21 days, donors 7–23) viability and gene expression experiments.

#### *4.3. Resazurin Viability Analysis*

Resazurin stock solution (AlamarBlue ®, Thermo Fisher, Waltham, MA, USA) was diluted 1:10 with serum-free cell culture medium. Slices were taken out of their hanging insert, washed once with phosphate-buffered saline (PBS) and transferred into a well of a 96-well plate filled with 300 µL resazurin solution. Incubation of slices was performed for 2 h at 37 ◦C and 5% CO2. After the incubation period, duplicates of 100 µL of the resazurin supernatant of each well were transferred to a fresh 96-well-plate. Fluorescence was determined on a plate reader (Synergy, BioTek, Winooski, VT, USA), applying excitation/emission wavelengths of 540/590 nm. Slices were placed back into their hanging inserts after viability determination.

#### *4.4. Confocal Laser Scanning Microscopy*

To validate results from the resazurin assay and to evaluate the spatial distribution of living and dead cells, additional live/dead determination using confocal laser scanning microscopy (CLSM) was performed on 13 slices of two donors (7 and 8) after three weeks of culture. Prior to the resazurin assay, two slices were heated to 60 ◦C for 30 or 60 min to decrease intra-tissue cell viability to a minimum and obtain a portion of low-viability slices in comparison to the control. Subsequently, a resazurin assay was performed on all slices. Afterward, the slices were analyzed optically via live/dead staining with fluorescein diacetate (FDA) and propidium iodide (PI; both Sigma Aldrich, St. Louis, MO, USA). Slices were taken out of their hanging inserts, washed twice with PBS and incubated in 6

µg/mL FDA in PBS solution (25 min), followed by a 0.1 mg/mL PI in PBS solution (3 min). After a washing step, slices were immediately transferred into the microscopes incubating chamber. Tissue imaging was carried out with a Nikon Scanning Confocal A1Rsi+ (Nikon, Tokyo, Japan) at 37 ◦C in DMEM without phenol red (Thermo Fisher, Waltham, MA, USA). Excitation/Emission wavelengths were set at 488/590 nm for PI and 485/514 nm for FDA. The scanning area was set at 2.82 mm<sup>2</sup> . Pictures were stitched with a 20% overlap and their corresponding volume was determined using Nikon Capture NX-D software (version 1.6.2, Nikon, Tokyo, Japan). Z-stacking was performed with 10 µm spacing. Z-stacks were rendered into a 3D image with living cells depicted in green and dead cells in red. Living cells were counted using ImageJ software (version 1.8.0. [34]). For comparison of both methods, resazurin assay viability data was correlated to the number of living cells per mm<sup>3</sup> sample using linear regression.

#### *4.5. RNA Extraction and Real-Time Quantitative PCR*

For RNA isolation, cartilage was carefully dissected from bony tissue, the resulting slices were soaked in RNAlater (Thermo Fisher, Waltham, MA, USA) and then stored at −80 ◦C overnight. The following day, the frozen tissue was transferred into liquid nitrogen and pulverized using a Biopulverizer (BioSpec, Bartlesville, OK, USA). Pulverized samples were transferred into a 90% TriReagent®, 10% 4-Bromo-2-chlorophenol solution (both Sigma-Aldrich, St. Louis, MO, USA) followed by centrifugation for 45 min at 13,000× *g*. The aqueous phase was collected, and nucleic acids were precipitated by the addition of an equal volume of ice-cold 70% isopropanol. After 30 min of incubation, precipitated nucleic acids were collected and resolved in RNA isolation buffer RLT (Qiagen, Hilden, Germany). Further purification was performed using a PicoPure™ Kit (Thermo Fisher, Waltham, MA, USA) according to the manufacturer's instructions. The integrity and purity of RNA were analyzed using an Agilent Bioanalyzer 2100 (Agilent, Palo Alto, CA, USA); RNA concentration was assessed with a NanoDrop 1000 spectrophotometer (Thermo Fisher, Waltham, MA, USA). RNA was reversely transcribed using a cDNA synthesis kit (iScript™, BioRad, Hercules, CA, USA). RT-qPCR was performed in triplicates in 96-well plates (Becton Dickinson, Franklin Lakes, NJ, USA) on a Mastercycler® ep gradient realplex (Eppendorf, Hamburg, Germany) using expression assays for TaqMan probes and primer sets (Thermo Fisher, Waltham, MA, USA; order no. in parentheses): collagen type II alpha 1 (*COL2A1*, Ss03373344\_g1), collagen type I alpha 1 (*COL1A1*, Ss003373341\_g1), aggrecan (*ACAN*, SS03373387\_S1), *Ki-67* (*MKI67*, qHsaCID0011882) and cartilage oligomeric matrix protein (*COMP*, Hs01572837\_g1). Expression analysis for matrix metalloproteinase 13 (*MMP13*, Ss033733279\_m1) was only performed on short-term cultured slices. Succinate dehydrogenase complex, subunit A (*SDHA*, Hs00188166\_m1) was used as reference gene. Marker gene expression is given as fold change compared to *SHDA* or control sample expression applying the efficiency corrected ∆∆-Ct method [35].

#### *4.6. Histological Analysis*

Slices of each donor were fixated overnight in 5% formaldehyde solution and subsequently decalcified for 21 days in Osteosoft® solution (Merck, Darmstadt, Germany). Afterward, slices were frozen in liquid nitrogen and subsequently cut into 4 µm thin sections using a CM19000 cryotome (Leica, Wetzlar, Germany). For Safranin-O staining, sections were stained for 30 min with 0.7% Safranin-O in 66% ethanol, counterstaining was performed with 0.2% Fast Green in 0.3% acetic acid (all Thermo Fisher, Waltham, MA, USA) for 1 min. To document ECM formation or loss, sections were mounted on glass slides; stainings were inspected using an AX 10 light microscope (Zeiss, Jena, Germany) and documented with a ProgRes® SpeedXT core 5 microscope-mounted camera system (JENOPTIK, Jena, Germany). The intensity of the Safranin-O staining is directly proportional to the glycosaminoglycan content of the tissue and was therefore analyzed employing a histomorphometrical approach as previously described [8]. Briefly, pictures were taken and all pixels in the areas of interest were valued in the RGB color mode with

a tool based on Xcode (Apple, Sunnyvale, CA, USA). When the red value (R) multiplied by 2 was higher than the sum of the green (G) and blue (B) values, the pixel was counted as red. The intensity of each red pixel was calculated as follows: intensity = 2 × *R*-value − *G*-value − *B*-value. Values of the intensity ranged between 1 and 508, and reporting images depicting the intensity distribution were created (see Supplemental Figure S3). The mean intensity (sum of intensities/area of interest) was calculated from each image.

#### *4.7. Statistical Analysis*

The significance level of log10-transformed data was determined with the independent two-sample *t*-test statistics of the Excel 2013 software package (Microsoft, Redmond, WA, USA). Normal distribution was checked applying the Anderson−Darling test, and equal variance of compared sample groups was tested applying the *f*-test. In all groups, signals were normally distributed. If the equal variance test was passed, student's *t*-test was used, if not Welch's *t*-test was applied. *p*-values < 0.05 were considered significant.

#### **5. Conclusions**

Native human osteochondral live slice explants represent a valid culture alternative to conventional punch explants. Its strengths lie in the greatly increased availability of tissue samples, as the thin nature of the slices allows for up to 50–100 pieces to be prepared from a single sample, the ease of fabrication and the simplicity of the culture in hanging inserts. This enables high-throughput screening of three-dimensional tissue processes and reactions in the native state that would otherwise require weeks of preparation in a tissue-based 3D culture or even in an animal model. Our results have shown that despite the stress of sharp dissection, the tissue fully retains its microarchitecture, and the cells retain their tissue-specific phenotype as well as their ability to properly respond to various stimuli. They also suggest that the short perfusion distance between cells and medium could also have a beneficial effect on the response strength of the cells to test stimuli, as the factors reliably reach cells embedded in the tissue. In addition, it was shown that the use of serum for cell expansion/survival is redundant in this culture form, which otherwise could mask adverse effects on tissue in the context of toxicity and degeneration analyses.

In summary, this production method is very well suited to produce native slice explants, which offer an interesting alternative to conventional explant cultures, especially in terms of pharmacological testing.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/ijms22126394/s1, Figure S1: *MMP13* gene expression analysis. Figure S2: technical details of custom-made microtome insert. Figure S3: examples of histomorphometrical analysis output.

**Author Contributions:** Conceptualization, J.S., T.D., L.K.S., U.S., A.A.K., C.R.; methodology, J.S., L.K.S. and T.D.; validation, J.S., L.K.S., A.A.K. and T.D.; formal analysis, T.D., S.H.-S. and C.R.; resources, C.R., U.S., D.K. and M.S.; data curation, J.S. and L.K.S.; writing—original draft preparation, J.S., L.K.S. and T.D.; writing—review and editing, J.S., L.K.S., T.D., C.R., A.A.K., S.H.-S., D.K. and M.S.; supervision, T.D., D.K. and M.S.; project administration, S.H.-S. and M.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Einstein Centre for Regenerative Therapies and Elke-Kröner-Fresenius Stiftung (Grant Number FKZ 2018\_T12).

**Institutional Review Board Statement:** The study was conducted in accordance with the Declaration of Helsinki, and the protocol was approved by the Ethics Committee of Charité-Universitätsmedizin Berlin (EA4/072/18).

**Informed Consent Statement:** Informed consent was obtained from all subjects involved in the study.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author. The data are not publicly available due to the use of patient material and the protection of personal data.

**Acknowledgments:** We thank our colleagues from Charité Research Workshop for crafting special instruments for slice culture manufacturing. We thank Christoph Müller and Anja Fleischmann for their assistance with tissue culture maintenance. We would also like to show our gratitude to the core unit cell harvest for organizing the logistics of the live tissue transport. Furthermore, we thank Henrik Mei from DRFZ for his excellent advice in the field of immunology of arthritic diseases.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Review* **Role of Physical Exercise and Nutraceuticals in Modulating Molecular Pathways of Osteoarthritis**

**Alessandro de Sire 1, \* , Nicola Marotta 1 , Cinzia Marinaro 1 , Claudio Curci 2 , Marco Invernizzi 3,4 and Antonio Ammendolia 1**

**Abstract:** Osteoarthritis (OA) is a painful and disabling disease that affects millions of patients. Its etiology is largely unknown, but it is most likely multifactorial. OA pathogenesis involves the catabolism of the cartilage extracellular matrix and is supported by inflammatory and oxidative signaling pathways and marked epigenetic changes. To delay OA progression, a wide range of exercise programs and naturally derived compounds have been suggested. This literature review aims to analyze the main signaling pathways and the evidence about the synergistic effects of these two interventions to counter OA. The converging nutrigenomic and physiogenomic intervention could slow down and reduce the complex pathological features of OA. This review provides a comprehensive picture of a possible signaling approach for targeting OA molecular pathways, initiation, and progression.

**Keywords:** exercise; physical activity; nutraceuticals; osteoarthritis; dietary supplements; inflammation; aging; inflammaging

#### **1. Introduction**

Osteoarthritis (OA) is one of the most common degenerative musculoskeletal disorders, characterized by a progressive loss of joint cartilage, synovial inflammation, formation of osteophytes, and subchondral bone remodeling [1–3]. This detrimental condition has a complex pathogenesis due to its multifactorial nature [4–6]. More specifically, the aberrant expression of degradative proteases or catabolic mediators might be induced in the chondrocytes, which contribute to cartilage erosion [7,8]. This imbalance between anabolic and catabolic processes might damage the structural integrity of the joint cartilage, resulting in stiffness, pain, and limited range of motion (ROM) in the later stages of OA [9,10]. Thus, subsequent loss of function, increased disability, lower performance in the activities of daily living (ADL), and reduction of health-related quality of life (HRQoL) are common findings in these patients [11].

In this scenario, an early diagnosis supported by the detailed understanding of the molecular pathways underpinning OA could help to develop tailored conservative therapeutic approaches aimed to avoid surgical and joint replacement treatments [12]. To date, several non-surgical treatments have been proposed in the last years for OA patients, including pharmacological treatments (e.g., acetaminophen, nonsteroidal anti-inflammatory drugs (NSAIDs), duloxetine, and opioids) [13], intra-articular injections with hyaluronic

**Citation:** de Sire, A.; Marotta, N.; Marinaro, C.; Curci, C.; Invernizzi, M.; Ammendolia, A. Role of Physical Exercise and Nutraceuticals in Modulating Molecular Pathways of Osteoarthritis. *Int. J. Mol. Sci.* **2021**, *22*, 5722. https://doi.org/10.3390/ ijms22115722

Academic Editor: Magali Cucchiarini

Received: 29 April 2021 Accepted: 26 May 2021 Published: 27 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

acid and glucocorticoids [14], focal muscle vibration [15], intra-articular oxygen-ozone therapy [16], radiofrequency ablation of genicular nerves [17], adipose-derived mesenchymal stem cell therapy [18], and platelet-rich plasma injections [19]. However, physical exercise is recommended by several guidelines as the first-line intervention in OA patients [13,20,21], which plays a crucial role in the prevention and treatment of the disabling sequelae of this severe chronic disease [22,23].

Nowadays, other alternative therapeutic interventions to treat OA have come to the fore, such as nutraceuticals, defined as substances that can be considered as a food or part of a food and provides medical or health benefits, including OA prevention and treatment [24,25]. In vitro and in vivo studies showed that epigenetic changes are triggered by micronutrients commonly present in diets (e.g., vitamins, carotenoids, and flavonoids) that might modulate OA mediator pathways [26–28]. Indeed, myoblasts and chondrocytes seem to share similar pathological targets and pathways and close anatomical location, suggesting the possible existence of a paracrine network [29]. In this context, an adequate prescription of physical exercise and nutritional supplementation might have a positive impact not only in terms of OA molecular pathways modulation, but also in terms of functioning and HRQoL improvement.

Therefore, in the present comprehensive review, we sought to describe the state-ofthe-art about the role that physical exercise and nutraceuticals might play in the complex management of OA, in terms of modulation of its molecular pathways.

#### **2. Osteoarthritis Molecular Pathways**

The pathogenic involvement in OA pathogenesis of pro-inflammatory cytokines released by chondrocytes and synoviocytes is well known and described in the scientific literature [7]. These cellular mediator patterns develop from cell signaling and gene expression pathways, which amplify the already altered cellular transduction, releasing additional inflammatory compounds and enzymes [30].

#### *2.1. Reactive Oxygen Species*

OA etiopathogenesis is influenced by several genetic and environmental factors not already fully explained. Recent studies have shown the involvement of oxidative stress and reactive oxygen species (ROS) in OA onset and progression [31]. ROS are unstable and highly reactive oxygen-containing free radicals combined with molecules to achieve chemical stability. Cells have developed antioxidant systems to scavenge ROS and maintain intracellular redox milieu. However, ROS are also key components of many physiological processes, and, at moderate concentrations, they act as indispensable second messengers. Their activity is based on the alterations of the cellular chemical environment, consisting of oxidative modification of proteins, influencing signal transduction, gene regulation, and cell cycling, in a complex process called redox biology [32]. The ability of cells to discriminate between the opposing effects of ROS is dependent on intensity, duration, and context of signaling and cellular redox status. When the cellular antioxidant capacity is insufficient to detoxify, ROS reacts with DNA, proteins, and lipids, disrupting their normal structure, impairing function, and leading to cytotoxicity in a process called oxidative stress [33].

Articular cartilage exists in relatively hypoxic conditions as a unique tissue thriving in a mechanically active environment. Despite living in a low O<sup>2</sup> environment, chondrocytes are characterized by abundant and active mitochondria that contribute to adenosine triphosphate (ATP) production [34]. Disruption of mitochondrial function, leading to increased levels of intracellular ROS, has been hypothesized to disrupt cartilage homeostasis and is considered as one of the main contributors of OA-related cartilage damage [35].

In physiological conditions, ROS are produced at low concentrations in chondrocytes mitochondria through oxidative phosphorylation and in cytoplasm by nicotinamide adenine dinucleotide phosphate (NADPH) oxidase. The overproduction of ROS observed in OA impairs mitochondrial functions through mtDNA damage, inducing chondrocyte

senescence and apoptosis, increasing cartilage degradation, and reducing matrix synthesis along with subchondral bone disfunction and synovial inflammation [36–38]. Evidence for ROS implication in cartilage degradation comes from the presence of lipid peroxidation products, nitrite, and nitrated products in the biological fluids in OA animal models [36]. On the contrary, antioxidant enzyme concentrations are decreased in OA patients, confirming the role that oxidative stress might play in OA pathogenesis [39].

Prolonged oxidative stress induces in the cartilaginous tissue the synthesis of large amounts of proteolytic enzymes, which in turn favors the shift towards catabolic reactions through several kinases such as p38 mitogen-activated protein kinase/nuclear factor kappa-light-chain-enhancer of activated B cells (p38MAPK/NF-κB) and activin receptorlike kinase 1 (ALK1) pathways [30]. In response to the proinflammatory stimuli, the overproduction of nitric oxide (NO) suppresses the cartilage matrix synthesis, enhances matrix metalloproteinases (MMPs) activity and induces chondrocyte apoptosis [40]. Taken together, all these mechanisms increase ROS concentrations leading to the membrane potential alterations that result in the release of cytochrome c and increase in Caspase-3 activity leading to apoptosis [41]. Lastly, oxidative stress-mediated inflammation is also responsible for the increased rate of hyaluronan degradation in synovial fluid [42].

On the other hand, chondrocytes respond to hypoxia and ROS damage, increasing the production of hypoxia-inducible factor-1 (HIF-1). HIF-1α can alleviate hypoxia-induced apoptosis, senescence, and matrix degradation in chondrocytes through mitophagy enhancing BCL2/adenovirus E1B 19-kDa-interacting protein 3 (BNIP3) expression [43].

#### *2.2. Cellular Apoptosis*

In the context of cellular autophagy, the phosphatidylinositol 3-kinase/protein kinase B (PI3K/AKT)/mechanistic target of rapamycin (mTOR) survival pathway plays a crucial role in OA pathogenesis. However, the role of Wnt/β-catenin and extracellular signal-regulated kinases (ERKs) pathway in OA pathogenesis is less elucidated [44]. In PI3K/mTOR signaling, AKT phosphorylated by PI3K modulates mTOR through a direct activation or an indirect block of one of its inhibitors. Activation of PI3K/mTOR pathway curbs the pro-apoptotic machinery resulting in enhanced cell survival through Caspase-3 inhibition [44,45]. Indeed, PI3K/mTOR is inactivated in IL-1b mediated chondrocyte apoptosis [46]. Alongside these mechanisms, Wnt proteins stand out through two characteristic pathways: β-catenin-independent pathway and β-catenin-dependent pathway [47]. In the independent process, the β-catenin undergoes phosphorylation, resulting in ubiquitination and consequent proteasomal degradation. On the other hand, in the β-catenin-dependent pathway, after the binding of Wnt ligands to the transmembrane frizzled (Fzd) receptor, the disheveled protein prevents the phosphorylation of β-catenin inducing the translocation into the nucleus where it stimulates the transcription of anti-apoptotic genes including c-Myc and cyclin D1 [48,49]. Lastly, recent studies seem to show that ERKs might be able to increase rather than decrease the levels of anabolic biomarkers in degenerative human chondrocyte [50–52].

#### *2.3. Pro-Inflammatory Signaling*

Alongside chondrocyte apoptosis, the scientific literature has recently focused on the mediation of inflammatory gene expression and inflammatory pathways role in OA pathogenesis. NF-κB is a protein complex that controls transcription of DNA and cytokine production [53]. In unstimulated cells, the NF-κB dimers are sequestered in the cytoplasm by a family of inhibitors called inhibitors of κB (IκBs) [54]. High Mobility Group Box 1 (HMGB1) is a non-histone DNA binding protein and is considered a damage-associated molecular pattern protein (DAMP) [55]. The binding of the extracellular ligand HMGB1 to the advanced glycation end product receptor (RAGE) has been described to activate IκB kinase (IKK), resulting in phosphorylation and degradation of IκBα [56].

Subsequently, p65 protein is released and phosphorylated for NF-κB heterodimer formation, and this complex is then translocated from the cytoplasm to the nucleus, inducing

the expression of several genes such as cyclooxygenase (COX), Matrix metalloproteinases (MMPs), and pro inflammatory cytokines [10].

This pathway is associated with mitogen-activated protein kinase (MAPK) signaling which constitutes a family of serine/threonine kinases (p38 MAPK; c-Jun amino-terminal kinase (JNK); and ERK1/2) characterized by a multilevel crosstalk and activated by a large array of inflammatory and stressful conditions [57,58]. In addition to the ERKs, the activated p38 MAPK can translocate into the nucleus upregulating kinases and transcription factors promoting cell autophagy in some cells while enhancing survival, growth, and differentiation in other cells [57]. Lastly, the phosphorylation of JNK via specific tyrosine and threonine residues is able to increase pro-inflammatory gene expression [58].

#### *2.4. Anti-Inflammatory Signaling*

Nuclear erythroid factor 2 (Nrf2) is a distress mediator involved in oxidative stress reduction [59]. It operates as a transcription factor binding to the antioxidant response elements (AREs) in the promoter region of antioxidant genes. In unstimulated chondrocytes, Nrf2 is kept quiescent through the association with kelch-like ECH-associated protein 1 (Keap1) in the cytosol, hence restricting its translocation into the nucleus to bind the AREs [60]. DJ-1 is a sensitive redox protein that serves as a stabilizer of Nrf2; in DJ-1 knockout animal models, the downregulation of Nrf2 could lead to an increase in oxidative stress. In the same context, heme oxygenase-1 (HO-1) is an Nrf2 downstream effector and both proteins are compensatory mediators that recognize pro-oxidative stressors and induce the production of antioxidant enzymes to neutralize the ROS [61]. Emerging evidence showed that the activation of DJ-1/Nrf2 signaling in chondrocytes is crucial to protecting these cells against oxidative insults, downregulating HMGB1 and NF-κB protein expression, and activating the PI3K/mTOR pathway [62].

In summary, the activation of NF kB, p38MAPK, and JNK is involved in the expression of several inflammatory genes that boost OA pathogenesis and progression. On the other hand, the upregulation of NRf2 could allow the expression of genes leading to a suppression of inflammation and inhibition of the activation of NF-κB. In this scenario, the PI3K/mTOR, Wnt, and ERK pathways appear to block cell apoptosis and promote proliferation.

#### **3. Physical Exercise as a Modulator of Osteoarthritis Molecular Pathways**

Physical exercise has been recognized as a safe, effective, and multifaceted therapeutic treatment to reduce pain and disability in OA patients [13,20,22,23,63].

The American College of Sports Medicine guidelines and EULAR recommend physical activity and exercise as positive and effective interventions on physical fitness as well as disease-specific and general outcomes in people with hip and knee OA. Moreover, these guidelines suggest that physical activity and exercise should be considered as integral parts of standard care [64,65].

Recommended exercise programs include land-based therapeutic exercise and water physiotherapy interventions [66,67]. Indeed, a recent systematic review confirmed that Pilates, aerobic, and strengthening exercise programs performed for 8–12 weeks, 3–5 sessions per week, with each session lasting 1 h, could be considered as an effective treatment in OA patients [68].

The exact protective mechanism of action of physical exercise on cartilage is not yet known, albeit several studies performed on animal models highlighted key molecules that could help define the best therapeutic exercise regimen.

Exercise modulates transcription in several metabolic pathways associated with extracellular matrix (ECM) biosynthesis and inflammation/immune responses in normal cartilage of rats undergoing treadmill walking (12 m/min, 45 min/day for 2, 5, or 15 days). More specifically, low-intensity exercises were able to regulate the different pathways involving PI3K-AKT, NF-κB, Ras, Rap, MAPK, cAMP, and Ptgs2. Moreover, exercise was able to suppress the expression of genes involved in ECM degradation, bone formation, and initiation of pro-inflammatory cascades, which are known to be upregulated in OA (Mmp9,

Mmp8, Igf1, ColIa1, Adamts3, Adamts14). Conversely, exercise can upregulate genes involved in ECM synthesis that are commonly downregulated in OA (Chrdl2, Tnfrsf11b, Timp4, Thbs2, Tgfb1, Mmp3, Il1r1, Il1r2, Cilp, and Bmp5) [29]. Taken together, these results suggest a crucial and multifaceted role of physical exercise in several molecular pathways involved in OA pathogenesis and progression.

#### *3.1. Chondroprotective Role of Physical Exercise*

OA animal models have been widely used in the research field to investigate the role of physical exercise in modulating molecular pathways involved in OA pathogenesis, highlighting a positive effect on cartilage preservation.

Iijima and colleagues showed that low-speed treadmill walking exercise (12 m/min for 30 min/day, 5 days/week performed for 2–4 weeks) prevented the progression of post-traumatic bone and cartilage lesions and increased BMP-2 and BMP-6 expression in the chondrocytes of the joint superficial zone of 24 male Wistar rats with induced damage to knee joints [69].

Assis et al. studied the effects of aerobic exercise performed with treadmill, training 3 days/week at 16 m/min for 50 min/day for 8 weeks, on an animal experimental model of knee OA. At the end of the treatment protocol, the exercise group had a better pattern of cartilage organization with less fibrillation and irregularities along the articular surface and a lower degenerative process, compared with controls. Moreover, animals in the exercise group had a lower chondrocyte nuclear or nucleolar expression of IL-1β, Caspase-3, and MMP-13, confirming the ability of aerobic exercise to downregulate proinflammatory, proteolytic, and apoptotic pathways [70]. Similarly, treadmill exercise with moderateintensity (18 m/min, 60 min/day, 5 days/week for 8 weeks) could have a chondroprotective effect in an OA animal model through the inhibition of NF-κB expression, resulting in a potent inhibition of MMP-13 gene expression [71].

Moderate-intensity exercise performed for only 4 weeks can also inhibit nuclear translocation of HDAC3/NF-κB complex, leading to the decreased expression of inflammatory proteins and cartilage protection in a rat model of OA. Similarly, in isolated primary rat chondrocytes mimicking OA-like pathologies, RGFP966, an HDAC3-specific inhibitor, could reduce IL-1β-induced inflammation and ROS production through the inhibition of nuclear translocation of NF-κB. Moreover, the same results were observed in a OA rat model, suggesting a role of RGFP966 in exerting exercise mediated chondroprotective effects [72]. Lastly, Castrogiovanni et al. demonstrated that a moderate-intensity physical activity protocol could lead to a decreased expression of OA-related biomarkers (IL-1, TNFα, MMP-13) and an increased expression of chondroprotective ones (IL-4, IL-10, and lubricin) in the synovium of an OA-induced rat model, highlighting the beneficial effects of exercise on cartilage preservation [73].

#### *3.2. Anti-Inflammatory and Anti-Apoptotic Role of Physical Exercise on Murine Models*

The anti-inflammatory effects of physical exercise in OA have been widely investigated on murine models. Here, a short review of the main findings showing the positive effects of physical exercise at different levels in OA damaged joints will be performed.

One hour treadmill activity showed an increase in Maresin-1 content, a strong antiinflammatory molecule, in intra-articular lavage fluid of MIA-induced rat OA. Maresin-1 was able to decrease MMP13, activate the PI3k/Akt pathway, and suppress the NF-κB p65 pathway in IL-1β-induced rat fibroblast-like synoviocytes in vitro, suggesting a pivotal role of Maresin-1 in exerting the anti-inflammatory activity of exercise [74].

Similarly, four weeks of moderate exercise in a monosodium iodoacetate (MIA) induced OA model determined IL-1β reduction and IL-4 increase in both serum and intra-articular lavage fluid, resulting in anti-inflammatory effect at cartilage tissue level. Furthermore, immunohistochemical assays showed that exercise markedly promoted the expression of autophagosome proteins LC3B, SQSTM1, and LC3II in the whole cartilage

tissue, suggesting a positive role of exercise in autophagosome's reaction to pathological stimuli [75].

Moderate intensity treadmill training in MIA-induced OA determined an increase in 15-hydroxyeicosatetraenoic acid (15-HETE) concentration in knee joint. 15-HETE is a well-known anti-inflammatory molecule and was also able to inhibit IL-1β-induced inflammation in primary chondrocytes and increase p-Akt levels in vitro. Moreover, 15- HETE injection in knee OA alleviated cartilage damage through the inhibition of MMP-13 expression and the concomitant increase in COL2 expression in joint cartilage tissue [76].

Other animal experimental models of OA included anterior cruciate ligament transection. A study performed on this specific animal model revealed that 4 months of moderate exercise determined the reduction of Caspase3 expression and concomitant overexpression of Hsp70, an antiapoptotic factor, in both superficial and deep zones of the knee medial compartment [77]. Similarly, in surgically-induced rat knee OA, low-speed treadmill exercise (12 m/min, 5 days/week for 4 weeks) increased pSmad-5, Id1 and BMP-2, BMP-4, BMP-6, and BMP receptor 2 expression in the superficial zone chondrocytes and suppressed cartilage degeneration [69].

In this scenario, it should be noted that physical exercise could provide beneficial effects in OA according to several factors, including adherence to exercise regimens, frequency of exercise, and actual loading of the symptomatic compartment [76,78]. Nam et al. [78] demonstrated that the extent of cartilage damage could play a key role in achieving the optimal effects of exercise. They conducted a study using a common treadmill exercise protocol (speed of 12 m/min for 45 min/day) after MIA-induction damage at knee level in rats, demonstrating that exercise significantly prevented the OA progression, albeit its efficacy appeared to be inversely related to the extent of cartilage damage. Transcriptome-wide gene expression analysis revealed that gentle treadmill activity started 1 day post–MIA OA induction significantly suppressed inflammation-associated genes through NF-κB network downregulation, with crucial effects on apoptosis and cell cycle (Cdkn1a and Bcl2), matrix breakdown (Mmp12 and Mmp14), and proinflammatory responses (IL15 and IL18) in murine models. On the other hand, a delayed intervention after grade 1 cartilage damage could be less effective in suppressing proinflammatory genes or upregulating matrix synthesis. These effects were mainly related to Sox9 suppression leading to the down-regulation of alkaline phosphatase, Cilp, Cilp2, and Mgp, which are crucial mediators required for correct matrix assembly [78].

#### *3.3. Beneficial Effects of Physical Exercise on Osteoarthritis Patients*

Evidence observed on rat and mouse OA models have provided further insights that have been translated to humans since the early disease stages. The beneficial effect of exercise has been already showed in subjects at high risk of developing OA and alterations in articular cartilage composition could be a good marker of early OA pathological modifications [79].

In a randomized trial, patients at a high risk of developing radiographic OA for partial medial meniscus resection were subjected to 4 months weight-bearing exercise protocol. Cartilage quality, comprising GAG, was evaluated through fixed-charge density tissue measurement by gadolinium-enhanced MRI (dGEMRIC). The exercise group showed an improvement in dGEMRIC, compared with controls (*p* = 0.036), suggesting that human cartilage has a potential to adapt to loading changes [80]. This adaptation consequent to exercises varies between different locations within the joint, and the largest beneficial effect is observed in the load-bearing cartilage, such as the lateral posterior cartilage [81].

dGEMRIC index changes were also investigated in the study of Munukka et al. The authors evaluated 87 postmenopausal women with mild knee OA performing leisuretime physical activity (LTPA). After a period of 12 months, a linear relationship between higher LTPA level and increased dGEMRIC index changes in the posterior region of the lateral and medial femoral cartilage, were observed. These results suggested that higher LTPA level is related to regional increases in estimated glycosaminoglycan content of

tibiofemoral cartilage [82]. Similarly, a randomized controlled trial conducted on 43 postmenopausal women with mild knee OA performing lower limb aquatic resistance training, showed a significantly reduction of T2-MRI scores in the posterior region of the medial femoral condyle after 4 months of physical activity. A decrease in T2 scores is suggestive of improved integrity and orientation of the collagen fibers and a decrease in hydration of articular cartilage [82].

Another promising technique to assess OA-related cartilage modifications is the microdialysis method. Helmark et al. obtained information about the effect of a single-leg knee-extension protocol on cartilage biomarkers and cytokines, both inside the joint as well as in the synovium, in a group of women affected by knee OA. In this RCT, over a period of three hours, a single bout of mechanical loading resulted in a significant decrease in cartilage oligomeric matrix proteins (COMP) and an increase in the anti-inflammatory mediator IL-10 in both intra-articular and peri-synovium spaces, compared with controls (*p* < 0.05). Significant increases were also registered for IL-6 and IL-8 in both groups, whereas TNF-α increased peri-synovially in the experimental group only [83].

Similarly, a pilot RCT performed by Hunt et al. demonstrated that a 10-week exercise protocol, consisting of hip abductors, quadriceps, and hamstrings muscle strengthening, performed on seventeen subjects with radiographically confirmed medial tibiofemoral OA, could lead to a reduction in serum of the cartilage degradation marker COMP levels, compared with controls (2.11 log U/L vs. 2.36 log U/L; *p* < 0.04). Moreover, a significant relationship emerged between dynamic measure of knee joint load and the ratio between two molecular markers increased in OA patient biological fluids (the urinary C-telopeptide of type II collagen/serum C-propeptide of type II procollagen ratio β = 1.11, 95% CI = 0.15, 2.07; *p* = 0.04) [84]. Lastly, the combined nutritional and exercise intervention also seem to act on cartilage biomarkers. A recent study demonstrated that an 18-month period aerobic walking and strength training in combination with an intensive diet determined the reduction of serum collagen type I (C1M) and II (C2M) in obese elderly patients affected by tibiofemoral OA [85].

#### *3.4. Physical Exercise as an Antioxidant Intervention*

The role of physical activity on oxidative stress and redox biology balance has recently obtained an emerging interest in the scientific literature. It is already known that physical activity might improve antioxidant defenses and lowers lipid peroxidation levels in both adults and aged individuals [86]. At cellular level, the beneficial role of physical exercise in OA comes through the increased expression of chondrogenic transcription factors SOX9 in circulating mesenchymal progenitors associated with autophagy. Autophagy regulates mitochondrial activity in stem cells to provide the best metabolic conditions, thus limiting ROS production and preventing metabolic stress and genome damage [87].

Furthermore, aerobic exercise modulates the chondrocyte response according to the balance between redox milieu and mechano-transduction mechanisms. Chondrocytes subjected to mechanical stimulation activate the NF-κB signaling pathway, which is considered a possible link between loading and chondrocytic responses to proinflammatory cytokines [88].

Nrf2 is a stress response protein in OA chondrocytes with anti-oxidative and antiapoptotic function and acts via ERK1/2/ELK1-P70S6K-P90RSK signaling axis in vitro [89]. Physical exercise could activate Nrf2 in response to the ROS increase, leading to the expression of antioxidant genes such as phase-II antioxidant enzymes such as glutathione and NADPH through Nrf2/Maf/ARE pathways activation [90].

Although researchers and clinicians have demonstrated a particular attention to the role of physical exercise antioxidant effects on OA, at present, evidence in literature about this issue is scarce. In MIA-induced OA rat model, 8 weeks of endurance exercise training (13 m/min, 50 min/day for 3 days/week) promoted free radicals generation that could stimulate both exercise adaptation and mitochondrial biogenesis. In this scenario, exercise promoted a significant increase in myeloperoxidase and superoxide dismutase activity

in the articular capsule [91]. Similarly, a two-group cross-sectional study showed that strengthening exercises could changes antioxidant status of systemic markers in knee OA patients. More specifically, ten obese women affected by knee OA, following an acute bout of isokinetic exercise, showed an immediate post-exercise rise of non-enzymatic antioxidant capacity of serum sustained by scavenging activity of 1,1-diphenyl-2-picrylhydrazyl. This finding suggests that the up-regulation of the body's antioxidant defense could be probably related to the mobilization of antioxidant reserves in plasma [92].

In the complex management of OA, physical exercise turned out to be effective also in synergy with other innovative therapies (i.e., mesenchymal cell implants or injections). In this scenario, autologous chondrocyte implants and bone marrow-derived mesenchymal cell implants or injections are innovative therapeutic interventions in OA focused on articular cartilage and subchondral bone damage treatment [93–95]. Studies in humans showed that exercise might enhance joint recruitment of bone marrow-derived mesenchymal stem cells and upregulates the expression of osteogenic and chondrogenic genes (Runx, MSx1, Sox9, COL2A1, ATG3), osteogenic microRNAs, and osteogenic growth factors (BMP2, BMP6) [87,96,97]. In experimental rodent models, physical exercise enhanced the osteogenic potential of bone marrow-derived mesenchymal stem cells and reduced their adipogenic potential. Moreover, exercise performed after stem cell implantation could enhance stem cell transplant viability [98–102]. Taken together, these results suggest a complex and multifaceted role of physical exercise on different antioxidant pathways corroborating the physiological basis supporting its efficacy in the treatment of OA.

#### *3.5. Challenges and Potential Controversies*

Despite its countless scientifically proven beneficial effects, physical exercise in certain conditions could be detrimental to articular cartilage health. Studies on human subjects reported that even moderate exercise, recommended as a treatment for OA, might increase markers of cartilage degradation in OA-affected joints [103,104]. Due to these controversies, the investigation of the imbalances in the molecular pathways involved in OA pathogenesis and progression could help in providing more insights about this controversial issue.

In an exercise-induced OA rat model, two weeks of 16 m/min treadmill activity showed that Ctnnb1, β-catenin, and Wnt-3a participate in the pro-inflammatory OA pathogenesis through an abnormal activation of the Wnt/β-catenin pathway due to excessive mechanical load [105]. Similarly a high-impact exercise protocol comprising flexion, extension, and compression of the limbs for two weeks resulted in IL-1β overexpression and IL-10 down-regulation in an advanced stage of OA induced in a rat model [106]. Siebelt et al. highlighted the impact of a 6-week treadmill intense exercise protocol in rats highlighting the loss of proteoglycan content in the exercised compared with sedentary OA joints induced by papain injection [104]. In surgically-induced rat knee OA, the same treadmill velocity was not able to modulate the effect of BMP pathway on OA pathological modifications, leading to the progression of morphological alterations [69]. Similarly, anterior cruciate ligament transected-animals running at a lower speed (18 m/min for 6 weeks) underwent advanced cartilage degradation, compared with the sedentary animals [107].

In humans affected by OA, Jayabalan et al. observed that 45 min of continuous walking resulted in a cumulative increase in COMP concentration, a marker of cartilage turnover, whereas interval walking was associated with COMP concentrations comparable to baseline. This study shed light about the possibility that incorporating resting periods in walking regimens may impact the potentially deleterious effects of longer continuous walking bouts on the knee joint [108].

In conclusion, the above-mentioned controversies might be due to the performance of high-intensity exercises for a long time. Indeed, they are not commonly recommended by the recent International Guidelines for frail OA patients [64,65].

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#### **4. Nutrigenomic: Role of Nutraceuticals on Osteoarthritis** ↓ and the NFκB signaling ↓ ↓ ↓ ↓ ↓

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Nutraceuticals are considered as dietary supplements including a concentrated form of a presumed bioactive substance, originally derived from a food, aimed to improve health status particularly in "active aging" [109]. In this scenario, nutraceuticals play a pivotal role on osteoarthritis pathogenesis with specific targets in modulating OA pathways (see Table 1). ↓ pathway by blocking IκB ↓ ↓ ↓ ↓ and the NFκB signaling pathway by blocking IκB → ↓ ↓ GlcN increases β ↑ ↑ ↑ ↓ ↓ ↓

**Table 1.**  However, they also exert their action in a high number of intracellular signaling and metabolic pathways [26]. PUFAs and their metabolites act as second messengers when intercalated in the cell membrane, inhibiting the expression of MMP-13 through the inactivation of the p38MAPK and NF-κB p65 pathway, and promoting the activation of the PI3k/Akt signaling [58,74].

> In the recent literature, the isolated administration of Chondroitin Sulfate has been shown to increase the transcription of the gene for Cartilage Oligomeric Matrix Protein (COMP) and decrease the expression of Toll-like receptors, thus inhibiting the Nfkb1 pathway in chondrocytes [112].

> Glucosamine (GlcN) is an amino monosaccharide component of glycosaminoglycan (GAG) chains, usually in the form of sulfate or hydrochloride salts derived from the chitin of the exoskeleton of crustaceans and from mushrooms [26]. Glucosamine promotes the proliferation of human osteoblasts modulating the mTOR pathway and enhancing β-catenin nuclear translocation with consequent activation of cell proliferation via cyclin D1 expression [49,121,137].

> Sulforaphane is an isothiocyanate derived from its glucosinolate precursor glucoraphanin, which is found in edible cruciferous vegetables, especially in broccoli. It can ameliorate ROS-induced oxidative stress and cartilage matrix degradation suppressing pro-inflammatory cytokines such as IL-6, TNF-α, and IL-17, inhibiting the JNK signaling and activating Keap1/Nrf2 pathway [133–135].

> Berberine is an isoquinoline alkaloid, produced by many plants, as Coptis japonica Makino, Coptis, Berberis petiolaris and B. vulgaris [138]. It modulates the PI3K/Akt and NF-κB pathways regulating cytoskeletal reorganization and dedifferentiation in chondrocytes, improving the progression of OA [111].

> Apigenin is a natural product belonging to the flavone class that is the aglycone of several naturally occurring glycosides. As an isolated compound from Cirsium japonicum

var. maackii, it blocks MMP3, MMP13, and COX-2 expression through Hif-2α inhibition via the NF-κB and JNK signaling pathways [110]. Therefore, downstream of the NF-κB and JNK signal expression might block the translation of the pro-inflammatory factor HIF2a.

Eupatilin is an O-methylated flavonoid present in Artemisia asiatica (Asteraceae), which suppresses the expression of pro-inflammatory genes in chondrocytes through the reduction of JNK phosphorylation, NF-κB, and MAPK activation pathway [28].

Other nutraceuticals that could have a role in modulating the OA pathways seem to be the green tea catechins (GTCs), which could protect against cartilage loss and reduce the progression of OA through the modulation of NF-κB inhibitor expression and targeting the PI3K/AKT pathway. Another main component of the green tea is the epigallo-catechin-3 gallate that might inhibit NF-κB nuclear translocation by blocking IkB degradation in human chondrocytes stimulated with IL-1β [122–124].

Genistein, an isoflavone from soy (Glycine max), inhibits the NF-κB pathway and the consequent expression of catabolic factors, while stimulating the expression of Ho-1, associated with the activation of Nrf-2 in human chondrocytes [118–120].

Furthermore, Jaceosidin, an extract of *Artemisia argyi*, showed to reduce the damage to OA cartilage by blocking the degradation of IκB and inhibiting the nuclear translocation of NF-κB [117].

Quercitin is a hydroxyflavonoid widely contained in the flowers, leaves, and fruits of different plants. It improved OA via dose-dependent effects on the Toll-like Receptor-4 (TRL-4)/NF-κB pathway in vivo, inducing a cytosolic down expression of TLR-4 [129].

Moreover, Wogonin is an O-methylated flavone present in *Scutellaria baicalensis* with a positive effect on OA, which may be related to the inhibition of pro-inflammatory genes expression via the NF-κB pathway [136].

Curcumin, the main polyphenol component from the roots of turmeric (*Curcuma longa*), improves the inflammatory pattern through NF-κB signaling inhibition and Nrf2/ARE signaling activation. In addition, curcumin inhibits cell autophagy via ERK1/2 signaling pathway activation [52,113–116].

Olive oil polyphenols (OOPs), as oleocanthal (OC), oleuropein (OP), tyrosol (TY), and hydroxytyrosol (HT), could modulate inflammatory and degenerative OA molecular patterns. OOPs inhibit pro-inflammatory genes expression through the suppression of NF-κB and MAPK activation, while enhancing Nrf-2 signaling [125–128].

Lastly, resveratrol, a phenol extracted from the skin of grapes, wines, mulberries, and peanuts, belongs to the phytoalexins that protect plants from microbial infections. Resveratrol increases cell proliferation via stimulation of Wnt signaling dependent pathway and might exert self-limiting mechanism of inflammation through PI3K/Akt and Nrf2 axis activation and concomitant NF-κB inhibition [47,86,139,140].

Therefore, nutraceuticals and physical exercise are involved in molecular pathways OA with apoptotic, pro-, or anti-inflammatory targets (see Figure 1 for further details).

**Figure 1.** Involvement of nutraceuticals and exercise in molecular pathways for apoptotic, pro-, or anti-inflammatory signaling.

#### **5. Conclusions**

In this comprehensive review, we described the main molecular pathways underpinning OA pathogenesis and progression, highlighting the different mechanisms of action and potential targets for innovative therapeutic interventions. Moreover, we described common targets of physical activity and nutraceuticals suggesting a possible therapeutical synergism of these two interventions. The decrease in ROS production upstream of physical exercise can guarantee a beneficial approach regardless of the nutraceutical chosen, whereas the mutual connections on a cellular mediator would suggest the composition of the compounds in an ad hoc logic.

In conclusion, despite the limitations related to the low characterizations of some molecular pathways, the current review provides intriguing insights regarding molecular pathophysiological mechanisms, innovative therapeutical target pathways, and synergistic interventions that could improve the complex framework of OA management.

— — **Author Contributions:** Study conceptualization, A.d.S.; methodology, A.d.S.; writing—original draft preparation, A.d.S., N.M., and C.M.; writing—review and editing, M.I. and A.A.; visualization, C.C.; supervision, A.d.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** Not applicable.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Review* **Vibrational Spectroscopy in Assessment of Early Osteoarthritis—A Narrative Review**

**Chen Yu 1,2,†, Bing Zhao 1,2,† , Yan Li 3 , Hengchang Zang 1,2,4,5 and Lian Li 1,2,4, \***

**Abstract:** Osteoarthritis (OA) is a degenerative disease, and there is currently no effective medicine to cure it. Early prevention and treatment can effectively reduce the pain of OA patients and save costs. Therefore, it is necessary to diagnose OA at an early stage. There are various diagnostic methods for OA, but the methods applied to early diagnosis are limited. Ordinary optical diagnosis is confined to the surface, while laboratory tests, such as rheumatoid factor inspection and physical arthritis checks, are too trivial or time-consuming. Evidently, there is an urgent need to develop a rapid nondestructive detection method for the early diagnosis of OA. Vibrational spectroscopy is a rapid and nondestructive technique that has attracted much attention. In this review, near-infrared (NIR), infrared, (IR) and Raman spectroscopy were introduced to show their potential in early OA diagnosis. The basic principles were discussed first, and then the research progress to date was discussed, as well as its limitations and the direction of development. Finally, all methods were compared, and vibrational spectroscopy was demonstrated that it could be used as a promising tool for early OA diagnosis. This review provides theoretical support for the application and development of vibrational spectroscopy technology in OA diagnosis, providing a new strategy for the nondestructive and rapid diagnosis of arthritis and promoting the development and clinical application of a component-based molecular spectrum detection technology.

**Keywords:** osteoarthritis; vibrational spectroscopy; near-infrared spectroscopy; infrared spectroscopy; Raman spectroscopy; early diagnosis

#### **1. Introduction**

Osteoarthritis (OA) is one of the major diseases affecting public health, and it is caused by both the mechanical and biological degradations of cartilage [1,2]. The 2017 Global Burden of Disease study shows that the annual number of OA patients is increasing, yet there remains no effective medicine to treat it [3]. OA is characterized by joint swelling, pain, and stiffness, causing physical pain to and placing a heavy financial burden on patients [4]. MacDonald et al. reported that the average age of people diagnosed with OA is 47.6, which is 7.7 years later than the onset of symptoms [5]. A Canadian study reported that patients with an OA score greater than or equal to 55 (severe disability) cost 3.5 times more than patients with an OA score of 15 (low severity) [6]. Most patients usually choose to see a doctor after they have obvious symptoms, and the general treatment of OA is often to address the corresponding symptoms, rather than to fundamentally cure the disease [7].

**Citation:** Yu, C.; Zhao, B.; Li, Y.; Zang, H.; Li, L. Vibrational Spectroscopy in Assessment of Early Osteoarthritis—A Narrative Review. *Int. J. Mol. Sci.* **2021**, *22*, 5235. https://doi.org/10.3390/ijms22105235

Academic Editor: Walter Herzog

Received: 1 April 2021 Accepted: 13 May 2021 Published: 15 May 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Many physiological and pathological changes may appear in the early stage of OA, which are the starting point for early diagnosis. OA is mainly caused by the inability of chondrocytes to maintain homeostasis between the synthesis and decomposition of intracellular and extracellular components, resulting in the inflammation of the synovium and joint capsule [8]. Moreover, various pathological changes may combine and worsen, leading to cartilage degeneration, subchondral osteosclerosis, angiogenesis, innervation, and other pathological phenomena. The extracellular matrix of chondrocytes is mainly composed of collagen, noncollagen proteins (glycosaminoglycan (GAG), proteoglycan (PG), cartilage oligomer matrix protein (COMP)), hyaluronic acid (HA), fibrin protein, follistatin-like protein 1 (FSTIL-1), glycoproteins, lipids, and water [9]. A number of these proteins are often used as biomarkers for the diagnosis of OA. GAG is the main component of the cartilage matrix. GAG in cartilage is cleaved by proteolytic enzymes, such as matrix metalloproteinases (MMPs) and aggregase, which decrease the expression level of GAG and lead to cartilage destruction. Therefore, GAG is often used as a marker of cartilage destruction in OA [10]. As an important component of synovial fluid (SF), HA can increase joint smoothness, thereby reducing cartilage wear, and it has been shown to be associated with the radiological progression of OA [3]. Subchondral bone can provide mechanical and nutritional support for cartilage. Small molecule diffusion experiments revealed that there is a direct molecular signaling pathway between cartilage and subchondral bone [11], indicating that changes in the microenvironment of subchondral bone may directly or indirectly affect cartilage metabolism. The quality and hardness of bone in patients with OA are decreased, making it more likely for some injuries to occur [12] and eventually leading to the production of osteophytes. In subchondral bone, markers such as the mineral-tomatrix ratio, mineral maturity/crystallinity, relative carbonate content, or relative tissue water content/porosity can provide signs of early OA [13]. The detection of changes in these markers is beneficial for an early diagnosis, which not only improves the treatment effect of OA for patients, but also greatly reduces treatment costs [14]. Considering the importance of an early diagnosis of OA, in this review, we aimed to summarize and analyze the vibrational spectroscopy used in early OA diagnosis to provide theoretical support for its application and development.

The diagnosis of OA primarily involves imaging and laboratory tests. The conventional imaging tests used include magnetic resonance imaging (MRI) [15–18], optical coherence tomography (OCT) [18–21], ultrasonic diagnosis [22–26], and X-ray [27–29]. Their specific applications are shown in Table 1. X-ray is the most accessible tool for assessing OA: it can show damage and other changes related to OA to confirm its severity according to different grading schemes, such as the Kellgren–Lawrence grading scheme [30]. MRI, which does not use radiation, is more expensive than X-ray but can provide better images of cartilage and other structures to detect early abnormalities in OA [16]. OCT is used to generate cross-sectional images of articular cartilage [19], and it can provide quantitative information on the state of articular cartilage, especially for OA caused by changes in collagen structure [31]. In terms of laboratory tests, the main method is joint aspiration, which comprises a needle being inserted into the joint, and fluid is extracted for further analysis to determine the stage of OA, which can help to rule out other forms of arthritis [32].

**Table 1.** Comparison table of application analysis of traditional imaging methods in relation to osteoarthritis (OA).

**Table 1.** *Cont*.

The methods mentioned above are commonly used in clinical practice but are often time-consuming, expensive, and even destructive. The development of rapid and nondestructive methods is urgent [19,29]. As a promising technique, vibrational spectroscopy has received increasing attention from researchers worldwide. A vibrational spectrum is generated by the vibration of molecules or atomic groups caused by electromagnetic radiation, and vibrational spectroscopy includes a variety of techniques, such as near-infrared (NIR), infrared (IR), and Raman spectroscopy [33]. As shown in Figure 1, NIR/IR/Raman spectra of SF, articular cartilage, and/or subchondral bone were collected to compare the spectral differences between healthy and OA-affected joints. The spectra of SF indicate that the content of protein increased and the content of carotenoid decreased in OA-affected joints [34]. Furthermore, the spectra of articular cartilage characterized changes in the contents of GAG and PG, as well as the orientation of collagen fibers in joints [35]. By analyzing the spectra of subchondral bone, it was concluded that mineralized components increased in OA-affected joints [36]. Extracting and analyzing effective spectral information can help to distinguish OA-affected joints from healthy ones, thereby improving the likelihood of early OA diagnosis. In this review, these new vibrational spectroscopy methods for OA diagnosis are introduced to provide theoretical support for their application and development in relation to the early diagnosis of OA.

**Figure 1.** Vibrational spectroscopy is used for early OA diagnosis by detecting changes in the composition and morphology of articular cartilage and changes in synovial fluid (SF) and subchondral bone. The figure depicts (**a**) a healthy joint and (**b**) an OA-affected joint.  The main biomarkers of OA in (**c**) SF and (**d**) articular cartilage are also shown. Vibrational spectroscopy can detect the changes of these components for early OA diagnosis.

#### **2. NIR Spectroscopy**

NIR spectroscopy reflects the overtones and/or combination bands of stretching and bending vibrations of C–H, N–H, and O–H bonds ranging from 12,500 to 4000 cm−<sup>1</sup> [37]. As shown in Figure 2, when a beam of NIR light with continuous wavelength irradiates to the sample, if the vibrational/rotational frequency of some groups in the sample is consistent with that of the NIR light, the molecule will absorb energy for energy-level transition, and the light at that wavelength is absorbed [38]. The sample can then be analyzed according to the transmitted or reflected light. NIR spectroscopy has been widely used in the agriculture [39–42], food [43–46], chemistry [47–49], and pharmaceutical fields [50–53] because it is fast, accurate, nondestructive and labor-saving. However, a limitation of this method is that the spectra are composed of wide overlapping bands [54,55], such that it is difficult to find specific peaks attributed to the complicated component. Chemometrics is a powerful tool for analyzing the NIR spectra and can improve the spectral resolution to help extract useful information from the spectra [56]. With the help of chemometrics, NIR spectroscopy has appeared in OA diagnostic research owing to its fast and nondestructive characteristics [57]. Although most of the work of NIR spectroscopy in OA diagnosis, including SF and tissue analysis, is still restricted to laboratory-scale research [58], it is becoming a useful tool for the diagnosis of OA and has good development prospects. −

**Figure 2.** SF is analyzed by NIR spectroscopy. The SF extracted from the joint capsule is dried and either membrane-formed or examined directly. The light source is divided into two beams by the splitter, which are then reflected onto two mirrors—one is a fixed mirror and the other is a moving mirror. After the light is reflected, it is recombined and sent to the sample, and then the change of the light beam after the reflection and absorption of the sample is detected. Finally, the spectral data are analyzed with the help of chemometrics.

#### *2.1. Application of NIR Spectroscopy on OA SF Analysis*