s2fieldsofstudy:
Biology
Materials Science

text:
Degradable magnesium implant-associated infections by bacterial biofilms induce robust localized and systemic inflammatory reactions in a mouse model

Biomaterial-associated Pseudomonas aeruginosa biofilm infections constitute a cascade of host immune reactions ultimately leading to implant failure. Due to the lack of relevant in vivo biofilm models, the majority of the studies report host immune responses to free-living or planktonic bacteria, while bacteria in clinical situations live more frequently as biofilm communities than as single cells. The present study investigated host immune responses to biomaterial-associated P. aeruginosa biofilms in a clinically relevant mouse model. Previously, we reported metallic magnesium, a prospective biodegradable implant, to be permissive for bacterial biofilm in vivo even though it exhibits antibacterial properties in vitro. Therefore, magnesium was employed as biomaterial to investigate in vivo biofilm formation and associated host immune responses by using two P. aeruginosa strains and two mouse strains. P. aeruginosa formed biofilm on subcutaneously implanted magnesium disks. Non-invasive in vivo imaging indicated transient inflammatory responses at control sites, whereas robust prolonged interferon-β (IFN-β) expression was observed from biofilm in a transgenic animal reporter. Furthermore, immunohistology and electron microscopic results showed that bacterial biofilms were located in 2D immediately on the implant surface and at a short distance in the adjacent tissue. These biofilms were surrounded by inflammatory cells (mainly polymorphonuclear cells) compared to the controls. Interestingly, even though the number of live bacteria in various organs remained below detectable levels, splenomegaly indicated systemic inflammatory processes. Overall, these findings confirmed the resistance of biofilm infections in vivo to potentially antibacterial properties of magnesium degradation products. In vivo imaging and histology indicated the induction of both local and systemic host inflammatory responses to P. aeruginosa biofilms. Even though the innate host immune defenses could not eliminate the local infection for up to two weeks, there was no apparent systemic bacteremia and all the animals investigated survived the infection.

Introduction
Applications of medical implants have increased manifold in humans for reasons such as organ loss due to chronic diseases and accidents [1]. Pseudomonas aeruginosa is the principal pathogen responsible for biofilm formation on medical implants [2,3]. Biofilms are the most precarious infections, since they are thousands of times more resistant to conventional antibiotics and host immune systems [4][5][6]. Biomaterial-associated biofilm infections are mostly treated by surgical removal of the implant from the infected tissue followed by the administration of antibiotics [7]. Such inevitable surgery together with antibiotic administration is extremely detrimental to patient health and the economy. Moreover, biofilm infections trigger unrestrained host immune reactions, which can cause immunosuppression, shock, transudation in organs, and defects in coagulation [8][9][10]. Innate and adaptive immune responses to various pathogens comprise accumulating phagocytic cells and inflammatory cytokines, among others interferons (IFNs) play a critical role [11,12]. However, the performance of host immune responses is compromised when most of the implant materials require a surgical procedure for their insertion into the body [13,14]. Initially, open wounds, body liquid-filled gaps between the implant and the surrounding tissue, structural, mechanical, and chemical properties of implants hinder immune cells from invading bacteria [15][16][17][18][19]. Later, bacteria secrete extracellular matrix material (ECM), which prevents phagocytic immune cells such as macrophages and neutrophilic granulocytes from engulfing the bacteria. The ECM also limits the diffusion rate of nutrients and metabolites, thereby reducing the bacterial metabolic activities. This impedes the efficacy of the proliferation-dependent antibiotics, thereby reducing the efficacy of various antibiotics [20][21][22]. To counter biofilm infections and augment the performance of implant materials, various strategies have been introduced. These include, metal alloy implants with antimicrobial properties, antibacterial as well as anti-adhesive nanoparticle coatings, and localized antimicrobial drug delivery systems on the implant surfaces [23,24]. Metallic magnesium as a biodegradable implant material is being highly investigated for clinical applications due to its in vitro bactericidal corrosion products and antibacterial properties, which can efficiently counter biomaterial-associated biofilm infections [25][26][27][28]. Magnesium alloys are presently under intense investigation as degradable, yet sturdy implant materials that could potentially be used for temporary applications to improve bone healing and avoid the long-term side effects associated with permanent metallic implants [29][30][31]. From our previous investigation as well as from the literature, magnesium and its alloys are reportedly susceptible to P. aeruginosa colonization and subsequent biofilm formation [32][33][34]. Limited information exists on the capacity of biomaterial-associated P. aeruginosa biofilms to escape the local innate immune defense and spread into the adjacent tissue or even systemically due to the lack of clinically relevant biofilm mouse models [35,36]. In addition, there is no in vivo information about the role of interferons against biofilm infections. Therefore, we employed in vivo P. aeruginosa biofilm formation on magnesium as a mouse model and report here P. aeruginosa biofilm repercussions directly on the implant interface, in the adjacent tissue, and systemically. We used a non-invasive in vivo imaging system to monitor the in vivo biofilm formation and interferon response to these infections [37]. Side by side, morphological, histological, and electron microscopic analyses were performed to determine biofilm effects on the host.

Preparation of magnesium samples and bacterial cultures
Disc-shaped magnesium samples (99.95% purity) having 5 mm diameter and 2 mm thickness were prepared and processed, as described previously [38]. Bioluminescent and wild type P. aeruginosa strains were streaked on LB agar plates and incubated for 18 h at 37°C [26,39]. For liquid cultures, isolates were cultured in BL broth medium and kept at 37°C with a shaking speed of 150 RPM until the desired OD 600 = 0.1 was achieved. The bacterial suspension was then immediately placed on crushed wet ice to inhibit further growth.

Surgical procedures and in vivo infections
Healthy 8 week old female wild type BALB/c mice were purchased (Harlan Winkelmann, Borchen, Germany) and IFN-β-reporter mice (IFN-β +/Δ β-luc ) on the BALB/c background (ifnb1 tm1.2Lien ) [40] were bred at the Helmholtz Centre for Infection Research, Braunschweig. The mice were kept in individually ventilated cages in a dedicated infection facility with water and food ad libitum. Before surgery, the animals were anesthetized with xylazine (4 mg Kg −1 ) and ketamine (10 mg Kg −1 ) and fur was removed at the site of surgery using a hair trimmer (Aesculap Suhl GmbH, Germany). Under sterile conditions in a laminar flow hood, 0.5 cm long incisions in the skin were made at three sites on the back of the mice using surgical scissors (Fine Science Tools GmbH, Heidelberg, Germany). A subcutaneous pouch was created from the incisions and 99.95% pure magnesium disks with 5 mm diameter and 2 mm thickness were inserted [26]. The wounds were closed by applying simple interrupted sutures (Ethicon Vicryl, Johnson & Johnson Medical GmbH). Immediately after implantation, logarithmically growing cultures of wild type P. aeruginosa (PAO1) or bioluminescent P. aeruginosa (PAO1 CTX::lux) at OD 600 = 0.1 were injected subcutaneously onto the surface of the implanted disks [41,42]. For monitoring the host interferon response, wild type P. aeruginosa (PAO1) were injected on subcutaneously implanted magnesium disks in the IFN-β reporter (IFN-β +/Δ β-luc ) BALB/c mice, which allowed tracking of the IFN-β gene induction [40]. D-luciferin (150 mg Kg −1 ) (Calipers) in phosphatebuffered saline (PBS) was injected intraperitoneally 15 min before examination by in vivo imaging. In vivo luminescence was recorded by using a Xenogen IVIS-200 optical imaging system (Caliper Life Sciences) equipped with an XGI-8 gas anesthesia unit (Caliper Life Sciences, Hopkinton, MA) using 2% isoflurane. The IFN-β-reporter mice were examined daily for a period of 2 weeks. The BALB/c mice were euthanized on the 8th day post infection and then the magnesium implants with adjacent tissue were removed and fixed in a buffered formalin solution according to standard procedures [32]. Animal experiments were performed with the permission of the Lower Saxony State Office for Consumer Protection and Food Safety (LAVES; Oldenburg, Germany) under approval number 33.42502/07-10.5.

Transmission electron microscopy (TEM)
The mice bearing infected implants were euthanized with CO 2 followed by cervical dislocation. The periimplant tissue was fixed in HEPES buffer containing 2.5% glutaraldehyde and 5% formaldehyde. The samples were washed with HEPES buffer and then incubated for one hour at room temperature in 1% aqueous osmium solution. The samples were dehydrated by serial incubation in acetone-water mixtures with increasing acetone concentrations (10%, 30%, and 50%) on ice, followed by a dehydration step in 70% acetone. Then, the samples were incubated overnight in 70% acetone and 2% uranyl acetate at 4°C. The following day, the dehydration process was continued in 90% and 100% acetone on ice and eventually in 100% acetone at room temperature. The samples were then embedded in epoxy resin, as previously described [43]. Ultrathin sections from these samples were prepared with a diamond knife, picked up with butvar-coated grids, counterstained with uranyl acetate and lead citrate, and examined in a transmission electron microscope (TEM910, Carl Zeiss, Oberkochen) set to an acceleration voltage of 80 kV. TEM images were digitally recorded at calibrated magnifications with a slow-scan CCD camera (ProScan, 1024 × 1024, Scheuring, Germany) using ITEM-Software (Olympus Soft Imaging Solutions, Münster, Germany). Brightness and contrast were adjusted with the help of an Adobe Photoshop version (Adobe).

Histological procedures
For histological analysis, the animals were sacrificed by CO 2 asphyxiation followed by cervical dislocation. The infected peri-implant tissues were isolated and fixed in 4% formalin for 3 d, as previously described in detail [44]. Briefly, the samples were dehydrated in a series of increasingly concentrated ethanol solutions and then embedded in paraffin. Histological slides of approximately 3 μm thickness were prepared from paraffin-embedded tissue samples. The sections were stained with hematoxylin and eosin (H&E) or periodic acid-schiff (PAS) staining according to standard laboratory procedures. Moreover, immunohistology staining for P. aeruginosa was performed with rabbitpolyclonal anti-Pseudomonas, (Biotrend, AP086) after heat-mediated antigen retrieval. The samples were evaluated histo-pathologically on microscope slides using a light microscope, randomized, and blinded to the experimental groups.
2.6. Tissue homogenization and colony-forming unit (CFU) assay One week after infection, the mice bearing biofilms were peacefully euthanized with CO 2 followed by cervical dislocation and their visceral cavities were opened. Visceral organs such as the liver, spleen, and gut were removed and their weights were measured. In parallel, the subcutaneously implanted magnesium disks inundated with biofilms and the adjacent tissue mass were isolated. Liver, gut, and peri-implant tissue were then placed in homogenization tubes containing 0.1% (v/v) X-100/PBS Triton solution. These tubes were then fixed in Polytron PT3000 homogenizer (Kinematica) and the homogenization process was carried out for 86 s. The explanted infected magnesium disks were directly submerged in PBS and vigorously vortexed for 5 min at maximum speed to disrupt the attached biofilm mass. The samples were analyzed for bacterial cell density (CFU/ml) by tenfold serial dilution and plating on LB plates at 37°C for 18 h.

Results
3.1. P. aeruginosa biofilm formation on magnesium implants in a mouse model A major challenge to establishing a biofilm mouse model is the rapid elimination of bacteria by the host immune system. To establish a mouse model, which supports biofilm infections for a prolonged period of time, we implanted subcutaneously three magnesium disks at selected locations in a mouse and then injected bioluminescent P. aeruginosa (PAO1) directly on each of these implants ( figure 1, d0). In agreement with our previous findings, the magnesium implants augmented the survival of the bacteria on their surfaces [32]. Bacteria formed biofilms on the magnesium and persisted for 2 weeks on the surface of these implants (figure 1, d0-d14). This experiment confirmed and reproduced our previous findings that magnesium promotes biofilm formation for an extended period of time. Previously, there was not a single mouse model which allowed biofilm formation on metallic implants with plain surfaces in the presence of an active host immune system. Therefore, we employed our system to investigate the distribution of biofilm infections and associated host immune responses.

Injected bacteria provoke a local interferon type-I immune response that can be visualized in transgenic mice by in vivo imaging
Type-I interferon response is triggered by various host cells and plays a critical role during the course of viral or bacterial infections. To investigate the interferon response to magnesium-based prolonged P. aeruginosa infections, we employed IFN-β reporter mice. In these transgenic mice, luciferase is expressed together with IFN-β, which is a virally and bacterially induced cytokine [40]. Magnesium implants were placed subcutaneously in the IFN-β reporter mice. The implanted magnesium disks were then infected with wild type P. aeruginosa (PAO1) and the IFN production was observed by in vivo imaging. Side by side, the IFN response measured from subcutaneously implanted sterile magnesium and P. aeruginosa injected into subcutaneous sham surgical pouches without implants served as controls. Immediately after infection, the IFN response was not within detectable limits of the system from the subcutaneous regions in mice carrying infected magnesium, sterile magnesium, or infected tissue (figure 2(A)d0, a-c). After 1 d, robust IFN-β-luciferase activities were detected from the sites that were infected either with P. aeruginosa only or in the presence of magnesium implants (figure 2(A), d1(a-c)). IFN-based luminescence was not observed from the sites with sterile magnesium implants during the entire course of the experiment (figure 2(A), d1b). In vivo expression of IFN-β induced by bacteria administered into the sham surgical pouches was detectable for 2 d and then disappeared. However, robust and prolonged IFN response detectable for 2 weeks was measured from the magnesium infected with P. aeruginosa ( figure 2(B), filled circles). Moreover, tissue necrosis was observed from these regions (pictures not shown). These results confirm our previous findings that magnesium implants support biofilm formation by P. aeruginosa, even in reporter mice. Prolonged IFN production and the resulting tissue necrosis confirm that P. aeruginosa biofilms provoke severe localized immune reactions.
3.3. P. aeruginosa biofilms cause intense accumulation of host immune cells A prolonged IFN response to P. aeruginosa biofilms provided evidence that these infections remain restricted at the site of infection. To confirm and explore the distribution of biofilms, implants after 1 week of infection were isolated together with adjacent tissue and subjected to electron microscopic analyses. Scanning electron microscopy (SEM) analysis revealed that P. aeruginosa encased in slimy material indicative of biofilms were clearly visible on the surface of the implants ( figure 3(A); asterisk symbols). In parallel, tissue pockets filled with bacteria were visible into the adjacent tissue ( figure 3(B); asterisk symbols). Bacterial intrusions into the tissue indicate the disseminating nature of biofilm infections ( figure 3(B)). Interestingly, this process occurred in the presence of large populations of host immune cells (figure 3(C); white arrows). TEM clearly indicated colonies of P. aeruginosa embedded deep into the tissue even in the presence of polymorphonuclear lecukocytes (PMN) (figures 3(D) and (E); white arrows). Electron transparent regions, a strong indication of biofilms, were clearly visible among the bacterial biofilms (figure 3(D); white arrows). P. aeruginosa was observed both intra-and extracellularly in PMN at the site of the biofilm formation (figure 3(F); asterisk symbols). This proves that the process of biofilm formation took place in the presence of active immune cells, which were indeed capable of engulfing the bacteria around the biofilms. Overall, it appeared that bacteria could utilize implant-tissue interfaces to establish biofilms and their distribution was limited to a short distance in the tissue where the spread of the infection was restricted by the host immune cells.

P. aeruginosa biofilms induce strong inflammatory responses in adjacent tissue
Host inflammatory response to bacterial invasion and foreign material is an expected phenomenon [45,46]. In contrast to planktonic cultures, biofilm bacteria persist longer within the host tissue even in the presence of an active immune system. To evaluate the intensity of the inflammatory reaction induced by P. aeruginosa biofilms, tissue sections were isolated from the mice bearing luminescent P. aeruginosa biofilms on magnesium implants after 1 week of infection and then subjected to H&E staining. For comparison, tissue sections from the sham surgical pouches infected with P. aeruginsoa and implanted with sterile magnesium were taken as controls. After 1 week, luminescent P. aeruginosa had established biofilms on the magnesium implants. Bacteria injected into a sham surgical pouch in the absence of implants disappeared within a few days. In an overview, the part of the tissue that housed the P. aeruginosa biofilms exhibited a high degree of inflammation with more obvious infiltration of severe neutrophilic inflammatory infiltrates surrounding the bacterially infected implants (figures 4(A) and (B)). Comparatively, the tissue portions that were infected with bacteria in the absence of implants or implanted with sterile magnesium elicited mild foreign-body reactions (figures 4(C)-(F)). These results strongly indicate that P. aeruginosa biofilm interactions with the tissue in the presence of magnesium provoke a high degree of inflammatory response.

Evidence of biofilm constituents in peri-implant tissue
Biofilm matrix is mainly composed of sugar polymers and proteins that can be visualized by PAS staining [47,48]. Even though electron microscopic analyses clearly revealed P. aeruginosa biofilms in the tissue, PAS staining was applied to further verify P. aeruginosa biofilms in the tissue near the infected magnesium implants. As expected, the PAS positive material indicative of the presence of biofilm matrix was detected in the tissue harboring infected magnesium implants (figures 5(A) and (B), black arrow). However, no PAS positive material could be detected near the infected areas in the absence of implants or in the tissue with sterile magnesium implants (figures 5(C)-(F)). The data also indicated that bacterial implant infections were not solely restricted to the implant surface. Instead, they expanded a short distance into the tissue and were walled off by the host immune cells.

Histological evidence of biofilm invasion into peri-implant tissue
Previous analyses confirm that tissues near the infected implants are filled with P. aeruginosa biofilms. To identify bacteria in the tissue, antibodies targeting P. aeruginosa were applied on the tissue sections isolated from the vicinity of the infected magnesium. Interestingly, a distinct region positive for Pseudomonas staining was visible in the tissue (figure 6(A), gray area marked by a black arrow). At high magnification, it could be seen that P. aeruginosa was dispersed within this particular area of biofilm formation ( figure 6(B), black arrow pointing to P. aeruginosa). The area positive for pseudomonas staining was surrounded by a high influx of inflammatory cells, which confirms biofilm resistance to the host immune cells ( figure 6(B), black arrow). Tissue bearing sterile magnesium or infected with bacteria only was negative for pseudomonas staining (figures 6(C)-(F)). Overall, various tissuestaining techniques confirmed that P. aeruginosa biofilms were not limited to the implant surface. Instead, they penetrate into the nearby tissue and trigger the induction of strong immune reactions (table 1).

Splenomegaly in mice with P. aeruginosainfected magnesium implants
Strong immunogenic and tissue inflammatory reactions associated with the presence of magnesiumbased P. aeruginosa biofilms convince us that these infections are not localized and can spread into the adjacent tissue. In the next step, the systemic effects of the biofilms were investigated by observing the liver, spleen and gut of mice bearing P. aeruginosa biofilms. These organs were then homogenized and the number of bacteria was determined. Bacterial CFUs could not be counted from the liver or spleen, except for the intestine, which seemed normal flora (table 2). The morphology of the intestine and liver of the mice bearing P. aeruginosa biofilms was normal and similar to uninfected animals. Surprisingly, the size as well as weight of the spleen isolated from the infected mice had increased compared to the uninfected mice ( figure 7(A) compare a with b). These results provide evidence that biomaterial-associated biofilm infections in the host affect systemic organs, particularly the spleen. Importantly, such a significant increase in the size and weight of the spleen is indicative of additional complications for patients suffering from biomaterialassociated infections.

Discussion
The results in this study confirm and refine previous results in that in vitro antibacterial properties of pure magnesium are not sufficient to prevent bacterial colonization of implants and instead apparently even promote prolonged bacterial survival under in vivo situations. Ours and some other studies have previously reported that in vitro antibacterial properties of magnesium are due to high alkalization. Therefore, one explanation for the absence of in vivo antimicrobial properties could be the absence of high pH in the peri-implant tissue controlled by the efficient bodybuffering system [26,49]. In addition, the development of conditioning film by the blood plasma or wound liquid creates a barrier for the dissemination of magnesium corrosion products, thereby allowing the colonization of bacteria [50,51]. Since hydrogen gas is a degradation product of magnesium corrosion, it cannot be excluded that the formation of gas pockets on the magnesium surface provides protection to the bacteria from early host cell invasion, and subsequently develop biofilms. Therefore, before clinical applications can be envisioned, it will be necessary to enhance the antibacterial properties of magnesium. Since magnesium alloys can be manufactured that have superior mechanical qualities, it could be envisioned to add antibacterial-acting metal alloys, such as copper or silver [52,53]. Since in this study no spontaneous infections of the magnesium alloy implants were observed, it may be sufficient to provide temporary protection for a limited time after surgery by antibiotic-releasing coatings until the wound has closed and the implant is protected by a fibrous capsule [54]. By the application of these strategies, the susceptibility of magnesium implants to biofilm infections could be reduced and it can be promoted for clinical applications.
P. aeruginosa is a significant pathogen responsible for implant failure in patients due to its biofilms. One important question about P. aeruginosa biofilms is whether the bacteria are localized on the implant surface, in the interface zone, or evade the tissue. Second, there is limited information about host immune responses to these biofilms. To answer these questions, the major challenge is the lack of clinically Figure 5. Evidence of biofilm ECM on the infected magnesium implants extending a short range into the peri-implant tissue. The tissue was fixed and subjected to PAS staining for histological examination by light microscopy. PAS positive staining of P. aeruginosa in the peri-implant tissue (A and B black arrow) propagated from the infected magnesium implants, PAS-stained tissue infected with bacteria in the absence of implants (C and D), and tissue bearing sterile magnesium implants (E and F). Scale bars on the left side correspond to 100 μm, scale bars on the right side to 26 μm. relevant animal models. Previous studies report that bacterial biofilms are mainly restricted on or near the surfaces of implant materials, because the mechanical and structural properties of these materials protect them from being engulfed by the host immune cells [55][56][57]. However, a recent study reported that S. epidermidis biofilms penetrated into the nearby tissues after initially growing on the implanted materials, resulting in robust inflammatory immune reaction [35]. However, in-depth characterization of biofilmmediated immune responses was not done in any of these studies. Here, we report for the first time biofilm-mediated immune responses in a highly sensitive reporter mouse expressing type-I interferon luciferase under transcriptional control of the IFN-ß promoter [40]. Our results indicate robust and persistent type-I Figure 6. P. aeruginosa infections originating from the medical implant surfaces extend into the adjacent tissue. As described in the legend of figure 1, the peri-implant tissue was fixed and subjected to P. aeruginosa-specific immuno staining. P. aeruginosa biofilms in peri-implant tissue (A and B black arrows), immuno-stained tissue infected with bacteria in the absence of implants (C and D), and tissue bearing sterile magnesium implants (E and F). Labeled structures are the same as those described in the legend of figure 3. Black arrows indicate regions with evidence of biofilm formation. Scale bars on the left side correspond to 100 μm, scale bars on the right side to 26 μm. Numbers indicate an arbitrary inflammatory score from 0 (no response observed in any of the three samples) to 4 (distinct inflammatory response in all three samples investigated). The data shown were compiled by an extensive analysis by an experienced histologist. However, at later stages, the bacterial number increased by biofilm formation resulted in IFN production. It also confirms that P. aeruginosa biofilms are recalcitrant to the immune system even in the presence of the IFN secreting active host cells. In addition, we restricted our research to the finding that strong IFN production is induced in response to P. aeruginosa biofilms. However, molecular mechanisms responsible for the activation of IFN in this mouse model need to be investigated. PMNs are powerful biocidal innate immune cells that can attack and occasionally even eliminate biomaterial-associated biofilms. The formation of P. aeruginosa biofilms in tissue together with PMNs indicates that implanted material provides initial niches for planktonic bacteria, which may otherwise be easily captured and killed by these cells in the absence of magnesium implants. At later stages, bacteria switch to biofilms and are strong enough to cope with immune cells. Consequently, bacterial biofilms from the implant surface penetrate into the nearby tissue and persist there as well. The detection of inflammatory cells, bacteria, and biofilm matrix components in the same tissue endorses that biofilms migrate from the implanted surface to adjacent tissue and are certainly resistant to the host immune system [58][59][60]. Biofilms in the tissue provide compelling evidence for the proliferating nature of these infections involving strong inflammatory reactions. Such a high influx of immune cells against biofilms could have the involvement of systemic organs, and therefore various body organs of the mice with biofilm-overgrown implants were observed. Interestingly, no systemic organ infiltration by bacteria could be detected by using the approaches described in this study. However, splenomegaly in mice with biofilms was observed. Splenomegaly is reported due to various reasons, but this is the first time that we have observed it in an animal suffering from biofilm infections [61,62]. It is possible that cytokines or migratory immune cells originating at the infected region could affect the inflammatory spleen response. Increases in the spleen size could nevertheless be indicative of systemic bacterial spread, perhaps at a level too low to detect by the methods applied here or it could be caused by the systemic distribution of inflammatory bacterial degradation products. The present study validated both localized and systemic inflammatory immune responses to bacterial biofilms established on magnesium implant surfaces. Thus, novel strategies need to be established to control such infections.

Conclusion
Persistent bacterial biofilms can form on degradable metallic magnesium implants in a small animal model. Luminescent-labeled bacteria allowed the monitoring of the course of the infection by in vivo imaging and the localized host immune response could be visualized by a transgenic mouse model. The resulting splenomegaly may be indicative of a low-level systemic spread of bacteria or bacterial products. In the absence of reliable conventional antibiotic treatments, novel strategies including antibiotic-acting implants may be envisioned to improve the performance of future magnesium-based implants. Figure 7. Prolonged P. aeruginosa biofilms on the magnesium implants lead to a significant increase in the size of the spleen. Magnesium implants were infected with growing cultures of P. aeruginosa or kept uninfected in individual mice. After 8 d, the mice were euthanized and their organs were removed and imaged with a light microscope (A). Spleen of mice bearing biofilm (A, a) and spleen of control mice (A, b). Changes in length (B) and weight (C) of the spleen isolated from the mice bearing infected or sterile magnesium implants.


extfieldsofstudy: Biology Medicine

text:
Influence of Mechanical Stimuli on Schwann Cell Biology

Schwann cells are the glial cells of the peripheral nervous system (PNS). They insulate axons by forming a specialized extension of plasma membrane called the myelin sheath. The formation of myelin is essential for the rapid saltatory propagation of action potentials and to maintain the integrity of axons. Although both axonal and extracellular matrix (ECM) signals are necessary for myelination to occur, the cellular and molecular mechanisms regulating myelination continue to be elucidated. Schwann cells in peripheral nerves are physiologically exposed to mechanical stresses (i.e., tensile, compressive and shear strains), occurring during development, adulthood and injuries. In addition, there is a growing body of evidences that Schwann cells are sensitive to the stiffness of their environment. In this review, we detail the mechanical constraints of Schwann cells and peripheral nerves. We explore the regulation of Schwann cell signaling pathways in response to mechanical stimulation. Finally, we provide a comprehensive overview of the experimental studies addressing the mechanobiology of Schwann cells. Understanding which mechanical properties can interfere with the cellular and molecular biology of Schwann cell during development, myelination and following injuries opens new insights in the regulation of PNS development and treatment approaches in peripheral neuropathies.

INTRODUCTION
Peripheral nerves are remarkable tissues of tremendous elasticity that propagate action potentials despite developmental growth (Vizoso and Young, 1948;Court et al., 2004;Simpson et al., 2013), stretches associated with movements of the limbs and mechanical compressions from daily activities Phillips et al., 2004). To resist mechanical stress, peripheral nerves are supported by three layers of connective tissues (i.e., epineurium, perineurium and endoneurium). In addition, each axon is individually wrapped by a large extension of modified plasma membrane called myelin sheath. Myelin appeared initially in hinged jaw fishes and was carried out in cartilaginous fishes. It contributed to the evolutionary success of larger vertebrates by allowing a faster and efficient conduction of nerves action potentials (Morell and Quarles, 1999), thus enabling faster predatory and escape maneuvers and the development of large and complex nervous systems (Zalc and Colman, 2000;Hartline and Colman, 2007;Zalc et al., 2008). In addition, to enhance the conduction of electric impulses and protect axons from mechanical stress damage, myelin producing cells also support the exchange of metabolites between the regional vasculature and the axon (Viader et al., 2011;Fünfschilling et al., 2012;Lee et al., 2012).
In the peripheral nervous system (PNS), Schwann cells (SCs) produce the myelin sheath. They are among the largest known cells, with 0.6 mm 2 of membrane on average (Rosenbluth, 2005). Understanding myelinating SC architecture can be challenging. To initiate myelination, SCs require an apicobasal polarity created by the axon (apical side) and the extracellular matrix (ECM; basal side). In addition, each myelin segment is flanked by nodes of Ranvier; thus creating a remarkable longitudinal polarity with internodal, juxtaparanodal and paranodal regions (Masaki, 2012; Figure 1). Yet, regardless of their gigantic myelin sheath and complex ultrastructure, SCs have a surprising plasticity. Upon injury to peripheral nerves, myelinated SCs can dedifferentiate and drive the regeneration of peripheral axons (Jessen and Mirsky, 2016;Boerboom et al., 2017). Outstanding reviews on SC development and the regulation of myelination in the PNS have recently been published and may be consulted for comprehensive characterization of SC biology (Kidd et al., 2013;Feltri et al., 2015;Monk et al., 2015;Salzer, 2015). In this review, we will focus on what the environmental constraints are for SCs, how SCs can sense and transduce mechanical signals and the experimental approaches that have started to unveil SC mechanobiology.

PERIPHERAL NERVES AND SCHWANN CELLS RESISTANCE TO MECHANICAL STRESSES
Peripheral nerves and SCs have adapted to sustain constant mechanical constraints. In peripheral nerves, myelinated fibers are surrounded by 6-15 layers of connective tissues (Sunderland, 1990) which shield SCs and axons from strains coming from the external environment (Lundborg and Rydevik, 1973;Rydevik et al., 1990;Wall et al., 1991Wall et al., , 1992Kwan et al., 1992;Brown et al., 1993). As previously mentioned, the connective tissue in peripheral nerves is constituted of three layers: the epineurium, the perineurium and the endoneurium. The epineurium constitutes a thick layer which surrounds the nerve and contributes to the resilience of the nerve (Peltonen et al., 2013). The perineurium is a strong layer of epithelium-like cells and collagen fibers (Thomas and Jones, 1967). Finally, the endoneurial ECM is found near the myelinated fibers, to protect them further from mechanical stresses. Interestingly, the endoneurial fluid is at a higher pressure than the epineurial fluid. An additional role for the perineurium could be to withstand the positive pressure from the endoneurium (Myers et al., 1978; Figure 1).
The stiffness and elasticity of peripheral nerves determine the mechanical cues that SCs are exposed to. They are partially determined by the connective tissue surrounding the nerves. Yet the elastic properties of peripheral nerves are not equal along their length. It is increased near articulations where nerves need to be more elastic, which reveals a more complex tissue architecture than anticipated (Phillips et al., 2004). For many years researchers have observed the undulating course of peripheral nerve fibers, also called spiral bands of Fontana FIGURE 1 | Schematic representation of the architecture of myelinated fibers in the peripheral nervous system (PNS). The three layers of connectives tissues are depicted and three myelinated fibers are enlarged. The first fiber (bottom) represents a rolled Schwann cell (SC). The second fiber (middle) shows a longitudinal section of a Schwann cell. The third fiber (top), is a view of Schwann cell unrolled. Axonal (orange •) and basal lamina (blue •) cell adhesion molecules (CAMs) and Linker of Nucleoskeleton and Cytoskeleton complex (LINC; purple •) are represented. Actin filaments are depicted with purple thin lines. SLI: Schmidt-Lanterman incisures. (Zachary et al., 1993). Bands of Fontana found in peripheral nerves disappear when nerves are stretched by 10% of their original length (Pourmand et al., 1994). Thus, bands of Fontana are thought to promote peripheral nerve elasticity. In support of this hypothesis, bands of Fontana are absent from nerves not subject to mechanical strain, such as newly regenerated nerves, intracranial nerves or spinal roots (Clarke and Bearn, 1972;Merolli et al., 2012). The mechanism by which bands of Fontana are formed is still unclear (Power et al., 2015).
SCs themselves adapted to endure external mechanical forces. Most of the SC myelin segment consists of compact myelin. Yet, compact myelin is interrupted by regions of uncompact myelin called Schmidt-Lanterman incisures (Small et al., 1987). Thus, it was proposed that incisures are a reservoir of membrane, allowing SCs to respond to a stretch of the myelin segment (Glees, 1943). While Schmidt-Lanterman incisures also have a role to play in SC intracellular transport (Balice-Gordon et al., 1998), it is important to point out that they are found only in the PNS, and their presence is conditional to the expression of P0, a major peripheral myelin protein (Yin et al., 2008(Yin et al., , 2015. SCs also contributes to the mechanical resistance of peripheral nerves by secreting the basal lamina. The basal lamina is an essential component of the SC ECM which is critical for SC development, surrounding each myelinated fiber Monk et al., 2015) and is an important component allowing regeneration of the peripheral nerves. The basal lamina organization remains unaffected after a severe mechanical compression on the nerve, or an acute axonal degeneration and is even further maintained after axonal death. As such, the basal lamina is able to be used as a guidance for axonal regrowth (Bunge et al., 1989). More recent methods have confirmed that the basal lamina provides a mechanical resistance to compression (Rosso et al., 2014). Autotypic junctions of SCs, formed between the spiral layers of myelin, have also been hinted to contribute to SC mechanical strength (Poliak et al., 2002). However, several models have shown that disruption of SC autotypic junctions impairs the propagation of action potentials but does not affect the appearance of the myelin sheath (Miyamoto et al., 2005;Denninger et al., 2015). Taken together, these studies suggest that relative elasticity or stiffness of the peripheral nerves influence the mechanical cues that the SCs are exposed to, while SC architecture and basal lamina integrity play key roles in the SC response to mechanical stress.

MECHANICAL INJURIES TO PERIPHERAL NERVES AND CONSEQUENCE TO SCHWANN CELL BIOLOGY
The pathological consequences of mechanical peripheral nerves injuries are evident, but their severity can vary. These injuries, were first classified by Seddon et al. (1943) ranked in three degrees: neurotmesis, axonotmesis and neuropraxia. Neurotmesis are dramatic injuries in which axons, connective tissues and basal lamina deposition are disconnected (Figure 2). They cause a complete loss of function. The recovery after neurotmesis is poor. Therapeutic approaches taking advantage of nerve grafts or nerve conduits have been attempted, but the success of such approaches have been mixed with variable improvements in functional recovery (Battiston et al., 2005). On the other hand, axonotmesis are severe compressions of the nerve or stretch-related injuries, and cause a disruption of axons and myelin sheaths with preservation of connective tissues and basal lamina (Figure 2). Distally to the point of trauma, axons degenerate and SCs demyelinate. Within 30 days after injury, peripheral nerves can remarkably recover, with FIGURE 2 | Myelinated fibers may be exposed to different stresses. (A) Tension strain can be caused by postures or movements. Under tension, the diameter of the myelinated fiber is also reduced, creating a transverse contraction strain. (B) Compression strain can be created by neighboring tissues (muscle, tendon, bone) or by environmental strains. External compression of peripheral nerves has been shown to alter myelin paranodal organization and can lead to demyelination (Dyck et al., 1990;Hodgson, 1993). Exposures to these stresses can caused various type of injuries, i.e., neuropraxia (1), in which myelin can be disrupted; axonotmesis (2), in which axons are disrupted; neurotmesis (3), in which the whole myelinated fiber, including its basal lamina are disrupted. axon regeneration and SC remyelination. Finally, in neuropraxia peripheral nerves endure more common and less severe trauma that can be caused by mechanical compression of the nerve or ischemia (Ochoa et al., 1971(Ochoa et al., , 1972. In neuropraxia, axon integrity is not primarily disrupted, but demyelination can occur (Figure 2). This injury affects the propagation of action potential, but is fully reversible, as the ''Saturday night palsy'' (Sunderland, 1990;Lee and Wolfe, 2000). Chronic compressions are another form of injury in peripheral nerves (Brown et al., 2011). They start with the inability of axons to transmit action potentials and as compression become more severe; demyelination follows (Rempel et al., 1999;Burnett and Zager, 2004). The effect of chronic nerve compression has been well characterized (Mackinnon et al., 1985;O'Brien et al., 1987;Dahlin et al., 1993;Pham et al., 2009). Chronic mechanical stimulation induces SC demyelination, as well as an increase in both SC apoptosis and proliferation, in absence of evident axonal injury (Gupta and Steward, 2003;Gupta et al., 2012). Similarly in vitro shear stress on primary SCs in the form of a laminar fluid flow is sufficient to reduce myelin protein gene expression and promote SC proliferation (Gupta and Steward, 2003;Gupta et al., 2005). Thus, low level mechanical stimuli may activate SC mitogenic pathways, independently from the regulation occurring between SCs and axons after axonal injury (Salzer and Bunge, 1980). It is also worth mentioning that genetic neuropathies could also be related to mechanical stresses. This appears to be the case with the hereditary neuropathy with liability to pressure palsies (HNPP). In this disease caused by the haploinsufficency of PMP22, patients are challenged daily by everyday activities. Benign mechanical pressures like crossing the legs can cause weakness or paralysis in the foot for several hours to several months (Earl et al., 1964;Horowitz et al., 2004;Li, 2015). The causes of HNPP are unclear as the exact role and functions of PMP22 are still being deciphered. Yet, a notable observation in HNPP myelinated fibers is the disruption of SCs autotypic junctions (Guo et al., 2014;Hu et al., 2016). Thus, it is possible that in SCs haploinsufficient for PMP22, the myelin is less resilient to mechanical stresses. This hypothesis is supported by atomic force microscopy work on myelinated fibers knock-out for PMP22 (Rosso et al., 2014). Thus, the response of SCs to mechanical stresses plays a role in both traumatic neuropathies and some genetic neuropathies.

EXTRACELLULAR AND INTRACELLULAR COMPONENTS OF SC MECHANOBIOLOGY
Mechanobiology refers to how physical and mechanical signals can be sensed by organs, tissues or cells and are converted in specific cellular responses. Many fundamental aspects of cell behavior depend on mechanobiology, including adhesion, spreading, migration, gene expression and cell-cell interactions in multiple cell-types (Jansen et al., 2015). Mechanobiology is based on the existence of two components: (1) mechanosensors, that allow a cell to sense mechanical signals provided by its environment; and (2) mechanotransducers, that allow a cell to transduce mechanical signals into biochemical signals. A large amount of mechanosensors and mechanotransducers from both intra and extracellular compartments have been identified over the last decades (Jansen et al., 2015). However these components are not relevant to all cells (e.g., ciliary bundles on hair cells in the sensory macula, glycocalix presence at the plasma membrane of epithelial cells, etc.; Jaalouk and Lammerding, 2009) and it is still unclear how mechanosensation and -transduction are orchestrated. In SCs, the identity of mechanotransducers and mechanosensors is starting to emerge. In the following section, we will review how physical signals can be transmitted in SCs through ECM, cell adhesion molecules (CAMs) and internal structures such as cortical cytoskeleton and the nucleus.

Extracellular Matrix
ECM acts as a physical support for cells to allow their adhesion and migration, and for tissue to protect their architecture from mechanical stresses. Its composition includes hundreds of proteins (i.e., elastin and collagen), glycoproteins and numerous proteoglycans. In addition, like epithelial cells, SCs secrete a basal lamina, an ECM component composed of laminin and collagen networks cross-linked with proteoglycans and nidogens, which links the extracellular environment to the plasma membrane of SCs . The mechanical properties of the ECM are defined mainly by elastic fibers and collagens fibers, which provide respectively resilience (elasticity) and tensile strength (resistance to tensile stress). In peripheral nerves, the proportion of elastic fibers in peripheral nerves is relatively small in comparison to collagen (Sunderland, 1965). In addition, collagen fibers are arranged longitudinally to allow some degree of axial stretch. Thus it was suggested that collagen fibers also contribute to the elastic properties of peripheral nerves (Tassler et al., 1994). Since all Schwann cells are surrounded by ECM, the most intuitive example of mechanotransduction is a transmission of signals through the ECM and the SC basal side. Yet, SCs also have an intimate relationship with axons on their apical side where biochemical signals from neurons are critical for SC migration, proliferation, survival, polarization, differentiation and gene expression (Monk et al., 2015;Salzer, 2015). For example, the myelinating fate of a SC is dictated by the amount of type III neuregulin-1 present on the surface of an axon (Taveggia et al., 2005). However, only membrane-bound type III neuregulin-1 promotes SC differentiation. In addition, a direct axonal contact is required for the deposition of a basal lamina by SCs (Bunge et al., 1982;Poitelon et al., 2015). These reports illustrate the functional need of SC to establish a direct physical contact with axons. Thus, SC fate depends on the formation of adhesion complexes between SCs with both axons and ECM. These adhesion complexes are primary sites of transduction for mechanical signals occurring in SCs and can be considered as mechanosensors.

Cell Adhesion Molecules
Major adhesion complexes are formed by CAMs and are implicated at various stages of SC development (Poliak and Peles, 2003;Feltri and Wrabetz, 2005 ; Figure 3). Regarding their role as mechanosensors, SC CAMs can be divided into two different groups based on their interaction with either the axon or the ECM (Figure 1). Each myelinating SC extends up to 1 mm in length along an axon, and keeps a minimal gap between the two at about 15 nm (Abe et al., 2004;Salzer et al., 2008). To maintain this proximity, numerous CAMs are expressed on both SCs and axons, including Caspr1, Caspr2, Contactin, Dystroglycan, Integrin β1, L1, MAG, Neurofascin 155 and 186, N-cadherin, Necl-2, Necl-4, N-CAM, NrCAM and Tag1 Berti et al., 2011;Colombelli et al., 2015). These CAMs are critical for the tight apposition and polarized organization of SCs, for review, see (Beirowski, 2013). Genetic studies have shown that ablation of several of these proteins leads to impairments of the interaction between the SC and the axon or its maintenance . In addition, stretch induced injuries could create a conduction block, caused presumably by the shear between axons and SCs at the nodes of Ranvier/paranodal region (Maxwell et al., 1991;Jou et al., 2000;Ichimura et al., 2005). The second group of CAMs, which distinctively interacts with the ECM, includes basal lamina receptors (i.e., integrin, G-protein coupled receptors and Dystroglycan). The role of these molecules has been well characterized during the last decades. SCs lacking basal lamina receptors experience detachment from the basal lamina and develop arrest or delay in the establishment of the interaction with an axon and its subsequent myelination Monk et al., 2015). Among these receptors, the role of integrin-mediated adhesions in mechanotransduction has been quite studied (Humphrey et al., 2014), and while some integrin-ECM interactions weaken when subjected to tension strain, often these adhesions strengthen or stabilize (Friedland et al., 2009;Litvinov et al., 2011). In particular, some β1 integrins have shown to strengthen under tension (Friedland et al., 2009;Kong et al., 2009). The behavior of SC α3β1, α6β1, α7β1 and α6β4 integrins under strain has not been defined yet (Berti et al., 2006), but is of interest as β1 integrins are essential for SC polarization and proper axonal ensheathment (Feltri et al., 2002).
Integrins are central to mechanosensation as they are connected both to the ECM and the actomyosin cytoskeleton, in clusters called focal adhesions. The structure of focal adhesions is complex and constitutes of numerous proteins. Thus, it is difficult to integrate the contribution of each one during mechanosensation. However, possible mechanically-induced stimulation of the focal adhesion complexes, could regulate important signaling pathways for SC biology, (i.e., downstream of FAK or ILK; Grove et al., 2007;Pereira et al., 2009;Grove and Brophy, 2014;Love et al., 2017), and stimulate the actin cytoskeleton in SCs. Indeed, Rac1 acts downstream of integrin β1 to drive actomyosin activity (Benninger et al., 2007;Nodari et al., 2007). The important role of actin in SC myelination was shown through the deletion of N-Wasp and Profilin1, regulators of actin polymerization (Jin et al., 2011;Novak et al., 2011;Montani et al., 2014). Yet, filaments of actin, linked to focal adhesions, also play a direct role in mechanosensation. The application of tensile force to cells increases their F-/G-actin ratio and promotes the formation of actomyosin filaments. In addition  .
to the actin assembly, activation of myosin II or myosin light chain kinase is critical for SC myelination (Fernandez-Valle et al., 1997;Melendez-Vasquez et al., 2004;Wang et al., 2008;Leitman et al., 2011;Montani et al., 2014). Thus, integrins can affect SC mechanobiology via their actions as mechanosensors and via changes in actomyosin activity.

Mechanotransduction Pathways
Although incompletely understood, the responses of the actomyosin cytoskeleton together with the activation of focal adhesion signaling induce several mechanotransducing pathways. The most notables are YAP and TAZ, two transcriptional activators of the Hippo pathway. Indeed, several works have shown that YAP can be activated through the Hippo pathway in SCs by Crb/Amolt proteins (Colciago et al., 2015;Fernando et al., 2016). In addition, mechanical stimulation through a signaling cascade involving FAK, Src, PI3K, JNK pathways (Codelia et al., 2014;Mohseni et al., 2014;Kim and Gumbiner, 2015;Elbediwy et al., 2016), or formation of actomyosin filaments and presence of F-actin (Dupont et al., 2011;Aragona et al., 2013) can regulate YAP and TAZ. Upon stimulation, YAP and TAZ are translocated to the nucleus to regulate gene transcription. In addition to regulating YAP and TAZ, F-/G-actin ratio also regulates the translocation of myocardin-related transcription factors (MRTFs) from the cytosol into the nucleus to activate serum response factor (SRF)dependent transcription (Olson and Nordheim, 2010). While YAP/TAZ and MRTF transcriptional targets overlap (Esnault et al., 2014), MRTFs have also been shown to be upstream of YAP and TAZ and to facilitate the YAP/TAZ response (Cui et al., 2015). Mechanotransduction by YAP and TAZ is critical to SC development and will be discussed in depth later in this review; however, the functions of MRTFs in the peculiar architecture of SC remain to be explored. The application of forces, including high density plating and mechanical compression, acts on the actin cytoskeleton and can lead to deformation of the nucleus and influence chromatin organization (Hernandez et al., 2016). The Linker of Nucleoskeleton and Cytoskeleton complex (LINC) binds the nuclear envelope and the actin cytoskeleton fulfilling a role of mechanotransducer between the cell inner membrane and its nucleus (Baarlink et al., 2013;Isermann and Lammerding, 2013;Plessner et al., 2015). Epigenetic modulators of chromatin structures, such as histone deacetylases, have been shown to be extremely important for SC myelination Jacob et al., 2011;Brügger et al., 2015;Wu et al., 2016), but the exact function of LINC in SC development is unclear.

Mechanosensitive Ion Channels
Although much emphasis has been placed on cell adhesion complexes as force sensors, mechanosensitive ion channels contribute both to mechanosensation and transduction. Over the past few years, a number of advances have been made through the identification of new ion channels, expressed in nearly all cell types, such as mechanosensitive potassium channels and Piezo ion channels (Coste et al., 2010;Ranade et al., 2015). They showed that conformational changes in the lipid bilayer of the plasma membrane, such as shear or membrane curvatures, can be converted directly in electric or biochemical signals (Brohawn et al., 2014). This is particularly of interest in the SC, which function is to wrap their membrane around an axon. Interestingly, the importance of membrane curvature during myelination has already been suggested in previous works, which have highlighted the role of lipids in plasma membrane (Ohler et al., 2004;Shaharabani et al., 2016) or N-Wasp (Novak et al., 2011). Considering that SCs express ion channels (Gray et al., 1984;Barres et al., 1990), the role of potential mechanosensitive ion channels in SC should be considered. In summary, SC mechanobiology can be influenced by both extracellular and intracellular cues. Particularly important regulators are the ECM, CAMs, YAP/TAZ signaling and the cortical cytoskeleton.

EXPERIMENTAL APPROACHES TO SC MECHANOBIOLOGY
Experimental approaches to study SC mechanobiology need to account for a variety of factors to model accurately the in vivo environment of peripheral nerves. During adult life, peripheral nerves are constantly subjected to mechanical stresses. In addition, the elasticity of peripheral nerves varies greatly along development, from 6 kPa after birth, when myelination is starting, to 50 kPa in adult nerves (Urbanski et al., 2016). This prompted the engineering community to investigate the response of SCs to their environment, in order to improve the design of nerve conduits used after injury (Gu et al., 2012;López-Fagundo et al., 2014). Isolated SCs grown on soft substrates (from 0.5 to 4 kPa) are rounded and present a low motility and a low proliferation rate (Gu et al., 2012;Poitelon et al., 2016). On stiffer substrates (≥40 kPa), SCs become bipolar and have an increased motility and proliferation (Gu et al., 2012;Poitelon et al., 2016). Surprisingly, SCs appear to be relatively insensitive to variations in stiff substrates, showing slight variations in morphology and no variations in the activation in YAP/TAZ mechanotransducers from 40 kPa to 4 MPa . However, when substrates are enriched with laminin, SC response is vastly different. On soft substrate coated with laminin (around 1 kPa), SCs are able to maintain a cytoskeletal architecture and form actomyosin filaments (López-Fagundo et al., 2014;Poitelon et al., 2016;Urbanski et al., 2016). In addition, the presence of laminin in stiff substrate (40 kPa) affects SC morphology, allowing them to spread, and activate YAP/TAZ mechanotransducers, as characterized by the transfer of YAP and TAZ in SC nucleus Urbanski et al., 2016). These data show that, in contrast to most anchoragedependant cells, SCs require laminin to respond to the stiffness of their substrate. The use of static two-dimensional monoculture to study the response of SCs to their environment remains limited. Peripheral nerves are continuously stimulated by stretched movements of the limbs and we do not know how compression, demyelination or loss of axons affects the stiffness of peripheral nerves.

Advances in Understanding SC Mechanobiology in Vivo
The first reports that studied YAP/TAZ in SCs in vivo confirmed that both mechanotransducers are active in SCs during axon-SC recognition, while SCs are myelinating and after completion of myelination Deng et al., 2017;Grove et al., 2017). During development, YAP/TAZ regulates SC proliferation, differentiation and myelination programs Poitelon et al., 2016;Deng et al., 2017;Grove et al., 2017). SCs ablated for both YAP/TAZ show an arrest of proliferation, fail to associate with an axon and do not initiate myelination Deng et al., 2017;Grove et al., 2017). The role of YAP/TAZ after myelination remains unclear as reports conflict on their requirements in adult SCs (Deng et al., 2017;Grove et al., 2017). Interestingly, YAP/TAZ regulates in concert several pathways including lipid or sterol biosynthesis in SCs but also integrins (i.e., integrin α6) and G-protein signaling (i.e., Gαs), suggesting respectively positive and negative feedback loops Deng et al., 2017). YAP and TAZ are functionally redundant, yet it appears that TAZ has a more prominent role in SC development Deng et al., 2017). Recent findings showed that after nerve injury TAZ levels are increased (Mindos et al., 2017). On the other hand, YAP levels remain stable in control animals and ablation of YAP alone does not affect peripheral nerve remyelination (Mindos et al., 2017). These data hint toward a specific role for TAZ in peripheral nerve remyelination. On the other hand, independent work demonstrated that YAP alone plays a role in the modulation of internodal length and positively regulates myelination and myelin elongation (Fernando et al., 2016). Interestingly, in a model deficient for the laminin receptor Dystroglycan, the amount of active YAP is reduced and internodes are shorter. However upon bone expansion, the peripheral nerves are stretched, the activity of YAP in SCs is increased and myelination appears to be improved (Fernando et al., 2016). These data suggest that physical stretch of the nerve could be a new therapeutic path to improve myelination. It is unclear if the physical stimulation provided by the stretch comes from CAMs at the axon interface, at the ECM interface or both. However, as myelin internodal length is known to grow together with limbs (Thomas and Young, 1949), it will be important to assess if the stretch affects also myelin thickness.

USING SC MECHANOBIOLOGY IN THERAPEUTIC APPROACHES FOR PERIPHERAL NEUROPATHIES
An exceptional quality of SCs is their ability to dedifferentiate from myelinated SCs to become repair SCs. After neurotmesis or axonotmesis, repair SCs are able to remyelinate newly regenerated axons, but most importantly, they provide the necessary tracks for axon guidance and support axonal regeneration. The function of repair SCs is highly considered in peripheral nerves injuries, but also in spinal cord injuries (Xu et al., 1995;Rutkowski et al., 2004;Oudega and Xu, 2006;Kanno et al., 2015). The non-invasive stimulations of SCs at the point of injury, or through the peripheral nerve in peripheral neuropathies, could be an approach to stimulate repair SCs to enhance recovery. The application of pulsed ultrasound on SCs in vitro appears to promote SC proliferation and prevent SC differentiation (Zhang et al., 2009;Tsuang et al., 2011). In vivo application of ultrasound after axonotmesis, also showed that SCs appeared to be affected, promoting an acceleration of peripheral nerve regeneration (Chang and Hsu, 2004;Raso et al., 2005;Jiang et al., 2016). Contrastingly, a most recent report showed that in SCs alone or in co-culture with neurons, ultrasonic stimulation promotes myelination signaling, with the increase of EGR2 levels in SCs (Yue et al., 2016). Thus it remains indispensable to precisely elucidate the influence of the ultrasound on SCs and peripheral nerves as well as the level and frequency of the ultrasound application. Another approach to stimulate SCs mechanically is the use of electromagnetic fields. Less studied, the stimulation of SCs by electromagnetic fields also opens interesting perspectives in peripheral nerve regeneration (Sisken et al., 1989;Kanje, 1992). Indeed, application of an electromagnetic field to cultured SCs promotes their proliferation and migration, presumably through YAP/TAZ (Colciago et al., 2015). Finally, physical therapy has also been considered as a therapeutic approach to stimulate tensile strain on peripheral nerves. Tensile stimulation, under a certain magnitude, appears to improve the regeneration of peripheral nerves (Bueno and Shah, 2008). However, much of this evidence is empirical and it is unclear how tension strain would stimulate axonal regrowth directly, or indirectly through SCs. In addition, the precise stresses required on a particular injured nerve are difficult to evaluate and could be more damaging. Thus patients could undergo such procedures only once the axon regeneration of nerve is completed (Suszynski et al., 2015).

CONCLUSION
SCs are poised to be the target of non-invasive mechanical therapies. Mechanically stimulating SCs could be of interest in traumatic neuropathies and demyelinating peripheral neuropathies, such as Charcot-Marie-Tooth disease, chronic inflammatory demyelinating polyradiculoneuropathy or Guillain-Barré syndrome. Yet, there is first a critical need to better understand SC mechanobiology. Determining the identity of the SC mechanosensors and mechanotransducers will define the signaling pathways and biochemical targets downstream of mechanical stress and help establish multidisciplinary approaches toward remyelination. Such research will also have the potential to be extended to strategies at reducing aberrant proliferation of SCs occurring in Schwannomas or neurofibromas.

AUTHOR CONTRIBUTIONS
SB and YP wrote the manuscript; SB, KLZ and YP revised the manuscript; YP created the figures. All authors approved the final version of the manuscript for publication.


extfieldsofstudy: Chemistry Medicine

text:
Levels of cyclic-AMP and cyclic-GMP in porcine oocyte-cumulus complexes and cumulus-free oocytes derived from small and middle follicles during the first 24-hour period of in vitro maturation

The objective of this study was to compare the cAMP and cGMP levels in cumulus-oocyte complexes (COCs) derived from the middle follicles (MFs, 3−6 mm in diameter) and small follicles (SFs, 1–3 mm in diameter) of pre-pubertal gilts during the first 24-h period of maturation in vitro (IVM). Both cAMP and cGMP levels in MF- and SF-derived oocytes did not change during this period. Although the cAMP levels increased in the COCs at 10 and 20 h after the start of IVM, the levels of cAMP were significantly higher in MF-derived COCs than in SF-derived COCs at 20 h after the start of IVM. On the other hand, the cGMP levels in COCs decreased to basal levels between 10 and 20 h after the start of the IVM, whereas cGMP levels were lower in SF-derived COCs than in MF-derived COCs during the first 10 h. The number of cumulus cells was larger in the MF-derived COCs than in the SF-derived COCs during the first 20-h period of IVM. The estimated cAMP level per cumulus cell at 10 h after the start of the IVM was higher in SF-derived COCs than in MF-derived COCs, whereas the estimated cGMP level per cumulus cell was no different between MF- and SF-derived COCs. From these results, we conclude that cAMP and cGMP levels in COCs, but not in oocytes, drastically change during the first 20-h period of IVM, and that both cAMP and cGMP levels significantly differ between MF- and SF-derived COCs.

S ince pre-pubertal pigs in many countries are commonly mostly slaughtered, ovaries from pre-pubertal females can be easily obtained for use as a source of gametes for embryo production in vitro. Techniques to produce embryos via in vitro maturation and fertilization (IVM-IVF) of the cumulus-oocyte complexes (COCs) derived from medium follicles (MFs; with 3-6 mm in diameter) of pre-pubertal ovaries have been well-developed [1,2]. However, the number of COCs from MFs is very limited on the surface of pre-pubertal gilt ovaries since small follicles (SFs) of less than 2 mm in diameter are the majority [3]. Furthermore, the meiotic and developmental competences of oocytes are significantly lower in SF-derived COCs than in MF-derived COCs [4][5][6]. Recently, we demonstrated that the follicular size significantly affects the meiotic and developmental competences. The capability of MF-and SF-derived oocytes to be fertilized and form pronuclei is similar only when mature oocytes containing the first polar body are selected following IVM [6]. Paracrine factors from the oocyte and gap junction-mediated communication seem to be involved in modulating large-scale chromatin structures during the final phase of oocyte maturation and these affect the acquisition of embryonic developmental competence in fully grown bovine oocytes [7]. We have also shown that autocrine factors from cumulus cells beneficially affect the meiotic and developmental competences of oocytes from SFs [8], and that a disruption of gap junction-mediated communication at around 20 h of IVM enhances the meiotic progression of SF-derived oocytes to the metaphase II stage [9]. However, it is still unknown why meiotic competence is lower in oocytes derived from SFs than in those derived from MFs; the differences in the resultant cells are also unknown. Additionally, both cAMP and cGMP are well-known secondary messengers that are involved in gap junction-mediated communications between the cumulus cells and the oocyte to affect oocyte resumption. The cAMP produced in the cumulus cells is transported into the oocyte through the gap junctions [10,11]. Increased intra-ooplasmic cAMP levels inhibit the activity of maturation-promoting factor (MPF) via the cAMP-dependent protein kinase pathway, which then consequently induces the arrest of oocytes at the germinal vesicle stage [12]. The cumulus cells produce cGMP though the natriuretic peptide precursor type C/natriuretic peptide receptor 2 signal pathway [13], which also negatively affects the rate of cAMP degeneration in oocytes by reducing the activity of phosphodiesterase-3 after being transported through the gap junctions, therefore preventing oocyte resumption during meiosis [13,14]. The activation of mitogen-activated protein kinases [15,16] by a gonadotropin-induced rise in cAMP levels [17,18] results in a reduction in the permeability of the gap junctions between the cumulus foot process and the oocyte. This occurs via the phosphorylation of proteins that are used to construct the gap junction and has been believed to induce meiotic resumption of the oocytes following a reduction in intra-ooplasmic cAMP and cGMP levels [15,16,19]. Therefore, the lower meiotic competence of oocytes derived from SFs than of those derived from MFs may be owing to changes in cAMP/cGMP reactions in the COCs/cumulus-free oocytes following gonadotropic stimulation until the breakdown of germinal vesicle during the first half of IVM. In the current study, we examined the cAMP and cGMP levels per COC or oocyte during the first 24-h period of IVM in MF-and SF-derived COCs before, during, and immediately after stimulation by dibutyryl-cAMP and gonadotropins.

In vitro maturation of COCs
Ovaries without any evidence of corpora lutea were collected from slaughtered commercial pre-pubertal gilts at a local abattoir and transported within 1 h to the laboratory at 32-35°C in a 0.9% (w/v) NaCl solution containing 75 µg/ml potassium penicillin G and 50 µg/ml streptomycin sulfate. The COCs were washed three times with the NaCl solution at room temperature and then were aspirated from either the SFs (1-3 mm in diameter) or MFs (3-6 mm in diameter) using a disposable 10 ml syringe and 18-gauge needle, separately stocked into a 50 ml centrifuge tube, and then washed three times with modified TL-HEPES-PVA. Only the COCs with a uniform ooplasm and a compact cumulus cell mass with at least 3 layers were then washed three times with mPOM. Approximately 30-40 COCs from SFs and MFs were separately cultured in 500 µl of mPOM supplemented with 10 IU/ml eCG, 10 IU/ml hCG, and 1 mM dibutyryl cyclic adenosine 3',5'-monophosphate (dbcAMP) in 4-well culture plates (Thermo Fisher Scientific, Roskilde, Denmark) for 20 h in an atmosphere of 5% CO 2 at 39°C [20,21]. The start of the IVM was considered to be when the COCs were first exposed to the gonadotropins and dbcAMP. The COCs were then washed three times with fresh IVM medium without gonadotropins and dbcAMP, and continued to be cultured in the medium for an additional 4 or 24 h.

Assessment of nuclear maturation
After IVM for 20 and 44 h, the cumulus cells surrounding the oocytes were removed by pipetting in a modified TL-HEPES-PVA medium containing 0.1% (w/v) hyaluronidase. Denuded oocytes were mounted on a glass slide and fixed in an ethanol:acetic acid (3:1, v/v) solution for 48-72 h at room temperature. The meiotic progression of the oocytes (111 oocytes from 3 replications and 393 oocytes from 5 replications at 20 and 44 h after the start of IVM, respectively) was evaluated under a phase contrast microscope at 200 × and 400 × magnifications after staining with 1% (w/v) orcein in 45% (v/v) acetic acid.

Measuring intracellular cAMP and cGMP contents
At 0, 10, 20, and 24 h of the IVM, the COCs from the SFs and MFs were separately transferred into pre-warmed TL-HEPES-PVA. The cumulus cells and zona pellucida were removed from some of the COCs by treating with 0.1% (w/v) hyaluronidase and 0.1% (w/v) protease, respectively, to obtain denuded oocytes. Thirty COCs or denuded oocytes were transferred into microtubes with 10 µl of TL-HEPES-PVA and stored at -80°C until they were assayed. The cAMP and cGMP levels in the COCs (1,680 and 930 COCs, respectively) and oocytes (1,590 and 1,080 oocytes, respectively) were measured using direct cAMP and cGMP enzyme immunoassay kits (Arbor Assays, Ann Arbor, MI, USA) and an iMark microplate reader (Bio-Rad, Hercules, CA, USA) according to the manufacturer's protocol. Briefly, 30 COCs and denuded oocytes were lysed in sample diluent (250 µl) and vortexed for 10 min at room temperature. After centrifugation at 2,200 × g for 10 min at 4°C, the supernatants (200 µl) were treated with acetylation reagent (10 µl) and then used for the assay. After adding a special plate primer (50 µl) to all wells of a microtiter plate, 50 µl of standards or samples were introduced into each well of the plate. Then, 25 µl of cAMP or cGMP peroxidase conjugate was added to all the wells. The binding reaction was initiated by adding an antibody for cAMP or cGMP (25 µl). After 2 h, the plate was washed and the substrate (100 µl) was added. After a short incubation for 30 min, the stop solution (50 µl) was added and then the intensity of the generated color was detected by using a microplate reader at 450 nm. The cAMP or cGMP concentrations (5-7 and 3-6 samples in each observation point, respectively) were calculated by using the Microplate Manager 6 software (Bio-Rad, Hercules, CA, USA).
To assess the status of the cumulus cells, the cAMP or cGMP levels per cumulus cell were estimated from the amount of cAMP/ cGMP in COCs or oocytes along with the number of cumulus cells in a COC as follows:

Assessment of the number of cumulus cells in a COC
At 0, 10, and 20 h of IVM, cumulus cells from thirty COCs were separated by treatment with 0.05% (w/v) trypsin in 0.5 mM EDTA solution for 10 min by pipetting. The number of cumulus cells per COC was calculated using a cell counting chamber. The experiments were replicated 5 times.

Statistical analysis
Data from 3-7 replicates were processed by a chi-square test (meiotic progression) or one-way ANOVA (amount of cAMP/ cGMP and the number of cumulus cells) using the GraphPad Prism 6 software (GraphPad Software, CA, USA). All data were expressed as mean ± SEM. Findings were considered statistically significant at P < 0.05 and when there was a significant effect in ANOVA. Values were compared by a Bonferroni-type Dunn post-hoc test.

Results
In total, 222 MF-and SF-derived oocytes (111 oocytes from each) from three replicated trials were assessed for meiotic stages at 20 h after the start of the IVM ( Table 1). The majority of the oocytes were at the GV-II stage, whereas the percentage was significantly higher in MF-derived (58.7 ± 1.9%) than in SF-derived oocytes (44.1 ± 2.6%). The incidence of oocytes at the GV-0 stage was significantly higher in SF-derived (23.4 ± 5.0%) than in MF-derived oocytes (8.8 ± 1.9%). When the meiotic progression of MF-(198 oocytes from 5 replications) and SF-derived (195 oocytes from 5 replications) oocytes was examined following IVM at 44 h, the percentage of oocytes that reached the metaphase-II stage was significantly higher (P < 0.05) in those from the MFs than in those from the SFs ( Table 2). The percentage of oocytes at the GV or metaphase-I stages was significantly higher in SF-derived than in MF-derived oocytes ( Table 2).
When the levels of cAMP in MF-and SF-derived COCs (1,680 total) and oocytes (1,590 total) were examined during the first 24 h after the start of IVM, the levels of MF-and SF-derived oocytes did not change (P > 0.05) throughout the period and did not differ significantly (P > 0.05) between the groups, although the cAMP levels in MF-derived oocytes displayed an upward trend around 10 h of IVM (Fig. 1B). However, cAMP levels in MF-and SF-derived COCs drastically changed just after the start of IVM, and were significantly higher at 10 and 20 h than at 0 and 24 h after the start of IVM (Fig. 1A). Furthermore, the levels of cAMP were significantly higher (P < 0.05) in MF-derived COCs (33.0 ± 0.5 fmol/COC) than in SF-derived COCs (28.4 ± 1.0 fmol/COC) at 20 h after the start of IVM. However, no significant differences (P > 0.05) in the cAMP levels were observed between MF-and SF-derived COCs at 0, 10, and 24 h after the start of IVM (Fig. 1A).
The levels of cGMP in MF-and SF-derived COCs (930 total) and the oocytes (1,080 total) were examined at 0, 10, 20, and 24 h after the start of IVM. As shown in Fig. 2B, the cGMP levels in the oocytes did not changed (P > 0.05) throughout the period and did not differ (P > 0.05) between MF-and SF-derived COCs. However, cGMP levels in both MF-and SF-derived COCs significantly decreased to a basal level between 10 and 20 h after the start of IVM ( Fig. 2A), whereas the cGMP levels at 0 and 10 h after the start of IVM were significantly lower (P < 0.05) in SF-derived COCs (41.7 ± 10.6 fmol/ COC and 24.8 ± 8.2 fmol/COC, respectively) than in MF-derived COCs (81.8 ± 4.5 fmol/COC and 64.8 ± 8.4 fmol/COC, respectively). There were no significant differences (P > 0.05) in cGMP levels at 20 and 24 h after the start of IVM regardless of the origin of the COCs.
Finally, the number of cumulus cells in MF-or SF-derived COCs was examined at 0, 10, and 20 h after the start of IVM (450 total COCs per group). Throughout this period, the number of cumulus cells was significantly larger (P < 0.05) in COCs derived from MFs than from SFs ( Table 3). As shown in Fig. 3A, the estimated cAMP level per cumulus cell was calculated by balancing the cAMP levels in a COC and an oocyte with the number of cumulus cells in a COC. The estimated cAMP levels increased significantly from 0 h (1.1 ± 0.2 amol/cell in MFs and 1.2 ± 0.5 amol/cell in SFs) to 10 h (7.2 ± 0.1 amol/cell in MFs and 11.1 ± 0.8 amol/cell in SFs) and 20 h (6.7 ± 0.4 amol/cell in MFs and 8.2 ± 0.5 amol/cell in SFs) after the start of IVM. Only at 10 h after the start of IVM, the estimated cAMP levels in a cumulus cell from a SF (11.1 ± 0.8 amol) were significantly higher than that from a MF (7.3 ± 0.1 amol). However, the estimated cGMP levels were no different (P > 0.05) between SF-derived (12.5 ± 3.6 amol/cell at 0 h and 5.3 ± 2.1 amol/cell at Table 1   10 h) and MF-derived (14.9 ± 4.6 amol/cell at 0 hour and 8.0 ± 3.4 amol/cell at 10 h) cumulus cells, whereas a downward trend (P > 0.05) was observed between 0 and 10 h after the start of IVM (Fig.  3B). The cGMP levels in the cumulus cells at 20 h were estimated to be at basal levels (-3.2 ± 1.4 amol/cell and -3.1 ± 1.8 amol/cell in MF-and SF-derived COCs, respectively).

Discussion
It is well known that the meiotic competence is significantly lower in COCs collected from SFs than in those collected from MFs [4,6,8,9]. In the present study, we observed that oocytes from MFs were at the metaphase-II stage and those from SFs were at the germinal vesicle or metaphase-I stage. The results presented here are consistent with previously published findings [6,8,9].
So far, some reports have shown different results regarding the dynamics of intracellular cAMP levels in MF-derived porcine oocytes during IVM [22−24]. Mattioli et al. [22] reported that the cAMP levels in COCs and oocytes did not change during the first 24 h (at 0, 3, 6, 12, and 24 h time points) after the start of IVM. On the other hand, Shimada and Terada [23] demonstrated that cAMP levels in COCs significantly increased at 4 h, peaked around 20 h, and then was maintained at that level until 48 h after the start of IVM (gonadotropins were supplemented through IVM for 48 h in this experiment). Furthermore, they also showed that the cAMP levels of oocytes increased between 0 and 4 h, peaked around 8 h,   and then started to decrease around 16 h before returning to basal levels around 32 h after the start of IVM. Additionally, Bagg et al. [24] reported that even in follicles with similar sizes in diameter, the dynamics in cAMP levels differed between pre-pubertal (no differences between 0 and 22 h of IVM) and post-pubertal gilts (significantly higher after 22 h of IVM). In the present study, we observed that the cAMP levels in MF-and SF-derived oocytes of pre-pubertal gilts did not change significantly during the first 24 h after the start of IVM. However, an upward trend in cAMP levels was observed in MF-derived oocytes after the start of IVM in the presence of dbcAMP (4.8 ± 0.5 fmol/oocyte at 0 h, 6.7 ± 0.7 fmol/ oocyte at 10 h, and 6.9 ± 1.6 fmol/oocyte at 20 h). Moreover, we observed that cAMP levels drastically increased in COCs in the presence of gonadotropins and dbcAMP and then decreased again in the absence of these factors. Therefore, changes in cAMP levels in cumulus cells may stimulate functions that consequently contribute to the meiotic resumption of the oocyte. However, the levels of cAMP were significantly lower in SF-derived COCs than in MF-derived COCs at 20 h after the start of IVM, even in the presence of dbcAMP. To our knowledge, this is the first study to report cAMP dynamics in SF-derived COCs and oocytes, although there was a paper that compared cAMP levels in COCs and oocytes derived from MFs of different sizes (3-8 mm in diameter) [25]. They showed that gap junctions between the oocyte and cumulus cells were maintained longer during IVM in COCs derived from follicles that were 3 mm in diameter instead of those that were 5-8 mm in diameter, and that the extent of cumulus expansion was also lower in COCs derived from smaller follicles [25]. Increased levels of cAMP in cumulus cells induced not only cumulus expansion [26,27], but also a disruption of the gap junctions in the COCs via the phosphorylation of connexin through the mitogen-activated protein kinase pathway [15,28]. Therefore, lower levels of cAMP in SFderived COCs during IVM may affect a lower meiotic competence in the oocytes due to insufficient cumulus expansion and a delayed disruption of gap junctions in the COCs. In fact, we also found here that the meiotic progression of the oocytes was significantly different between the MF-and SF-derived COCs, since more oocytes were at the GV-II stage in MF-derived COCs and at the GV-0 stage in SF-derived COCs at 20 h after the start of IVM when supplemented with gonadotropins and dbcAMP. In a previous study [21], we have also shown that the incidence of oocytes at the GV-II stage increased when MF-derived COCs were cultured with gonadotropins and dbcAMP. A difference in the reaction against gonadotropins and dbcAMP between MF-and SF-derived COCs may be the cause of the observed disparity in cAMP levels and the meiotic progression during the first 20-h period of IVM. Recently, we have also reported that removing cumulus cells from SF-derived COCs 20 h after the start of IVM significantly improved the meiotic competence of the oocytes [9]. Therefore, it may be important to achieve a suitable reaction for SF-derived COCs against gonadotropins and dbcAMP to induce a well-timed disruption of gap junctions between the cumulus cells and the oocyte.
Furthermore, in the present study, we examined the cGMP levels in MF-and SF-derived COCs and oocytes at 0, 10, 20, and 24 h after the start of IVM. To our knowledge, this is also the first paper to report cGMP dynamics in MF-and SF-derived COCs and oocytes during IVM in pigs. We found that levels of cGMP significantly decreased to basal levels in both MF-and SF-derived COCs between 10 and 20 h after the start of IVM in the presence of gonadotropins and dbcAMP. After a hCG injection in vivo, the levels of cGMP have been shown to decrease in mouse ovaries [29] and oocytes [14]. Previous studies have also shown that cGMP levels were significantly reduced in isolated follicles following the addition of luteinizing hormone to the medium [14,16,29]. These previous reports in mice are consistent with our current results.
We have also observed that levels of cGMP were significantly lower in SF-derived COCs than in MF-derived COCs at 0 and 10 h after the start of IVM, although the cGMP levels in oocytes did not differ. Production of cGMP via the NPPC/NPR2 pathway has been shown to be accelerated by oocyte-secreting factors and 17β-estradiol [13,30]. Since intra-follicular 17β-estradiol concentration increases with the growth of the follicle [31], this factor may be affecting the observed difference in cGMP levels between the MF-and SF-derived COCs.
The present study also demonstrated that the number of cumulus cells was significantly lower in SF-derived COCs than in MF-derived COCs through IVM. These results are consistent with previous reports  [5,32]. When the estimated levels of cAMP per cumulus cell were calculated from the current results, the overall dynamics reflected the cAMP levels in COCs. However, no differences between the cAMP levels in MF-and SF-derived cumulus cells were observed within 10 h after the start of the IVM. Furthermore, the estimated cGMP levels did not differ significantly through IVM between the MF-and SF-derived cumulus cells. These results suggest that the total number of cumulus cells per COC may reflect the total cAMP and cGMP levels in COCs as well as the meiotic progress of the oocytes during IVM. Further research is required to investigate whether an increase in the number of cumulus cells surrounding the SF-derived oocyte before or during IVM will improve the meiotic and developmental competence of the oocytes. Additionally, it is generally believed that cGMP plays a role in COCs to inhibit the decomposition of cAMP by inhibiting phosphodiesterase-3 activity [13,16], which consequently maintains meiotic arrest at the germinal vesicle stage. On the other hand, the cGMP/ protein kinase G pathway is associated with lipid metabolism in adipocytes [33,34]. Since lipid metabolism in cumulus cells affects the viability and meiotic competence of bovine oocytes [35], the role of cGMP in the meiotic and developmental competences of COCs should be further investigated.
In conclusion, both the levels of cAMP and cGMP in COCs drastically change during the first 20-h period of IVM, and these levels differ significantly between MF-and SF-derived COCs. These differences may significantly affect the meiotic competence of MF-and SF-derived oocytes.