Synthetic niche to modulate regenerative potential of MSCs and enhance skeletal muscle regeneration

Accepted Manuscript Synthetic Niche to Modulate Regenerative Potential of MSCs and Enhance Skeletal Muscle Regeneration Matthias Pumberger, Taimoor H. Qazi, M. Christine Ehrentraut, Martin Textor, Janina Kueper, Gisela Stoltenburg-Didinger, Tobias Winkler, Philipp v. Roth, Simon Reinke, Cristina Borselli, Carsten Perka, David J. Mooney, Georg N. Duda, Sven Geißler PII:

S0142-9612(16)30176-4

DOI:

10.1016/j.biomaterials.2016.05.009

Reference:

JBMT 17491

To appear in:

Biomaterials

Received Date: 8 March 2016 Revised Date:

2 May 2016

Accepted Date: 4 May 2016

Please cite this article as: Pumberger M, Qazi TH, Ehrentraut MC, Textor M, Kueper J, StoltenburgDidinger G, Winkler T, Roth Pv, Reinke S, Borselli C, Perka C, Mooney DJ, Duda GN, Geißler S, Synthetic Niche to Modulate Regenerative Potential of MSCs and Enhance Skeletal Muscle Regeneration, Biomaterials (2016), doi: 10.1016/j.biomaterials.2016.05.009. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Title: Synthetic Niche to Modulate Regenerative Potential of MSCs and Enhance Skeletal Muscle Regeneration

† Authors contributed equally as first authors ‡ Authors contributed equally as senior authors * Corresponding author

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Affiliations:

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Matthias Pumberger1,2,3 †, Taimoor H. Qazi1,2 †, M. Christine Ehrentraut1, Martin Textor1,2,3, Janina Kueper1, Gisela Stoltenburg-Didinger4, Tobias Winkler1,2,3, Philipp v. Roth1,2,3, Simon Reinke1,2,3, Cristina Borselli2,5, Carsten Perka1,2,3, David J. Mooney2,5, Georg N. Duda 1,2,3,5‡, Sven Geißler1,2,3‡ *

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Julius Wolff Institute, Charité – Universitätsmedizin Berlin, Augustenburger Platz 1, 13353 Berlin, Germany

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Berlin-Brandenburg Center for Regenerative Therapies & Berlin-Brandenburg School for Regenerative Therapies, Charité – Universitätsmedizin Berlin, Augustenburger Platz 1, 13353 Berlin, Germany

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Center for Musculoskeletal Surgery, Charité – Universitätsmedizin Berlin, Augustenburger Platz 1, 13353 Berlin, Germany 4

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Institute of Cell Biology and Neurobiology, Charité – Universitätsmedizin Berlin, Charitéplatz 1, 10117 Berlin, Germany

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Wyss Institute for Biologically Inspired Engineering, Harvard University, 3 Blackfan Cir., Boston, MA 02115 (USA); John A. Paulson School of Engineering and Applied Sciences, Harvard University, 29 Oxford St., Cambridge, MA 02138 (USA).

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*Correspondence: E-mail: [email protected], Fax: +49-30-450559969, Telephone: +49-30-450659539

Key words: Engineered cell microenvironments, Biomaterial based stem cell niche, Trophic factor secretion, Muscle regeneration

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Abstract: Severe injury to the skeletal muscle often results in the formation of scar tissue, leading to a decline in functional performance. Traditionally, tissue engineering strategies for muscle

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repair have focused on substrates that promote myogenic differentiation of transplanted cells. In the current study, the reported data indicates that mesenchymal stromal cells (MSCs) transplanted via porous alginate cryogels promote muscle regeneration by secreting bioactive

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factors that profoundly influence the function of muscle progenitor cells. These cellular functions, which include heightened resistance of muscle progenitor cells to apoptosis,

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migration to site of injury, and prevention of premature differentiation are highly desirable in the healing cascade after acute muscle trauma. Furthermore, stimulation of MSCs with recombinant growth factors IGF-1 and VEGF165 was found to significantly enhance their paracrine effects on muscle progenitor cells. Multifunctional alginate cryogels were then utilized as synthetic niches that facilitate local stimulation of seeded MSCs by providing a

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sustained release of growth factors. In a clinically relevant injury model, the modulation of MSC paracrine signaling via engineered niches significantly improved muscle function by remodeling scar tissue and promoting the formation of new myofibers, outperforming

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standalone cell or growth factor delivery.

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1. Introduction

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Muscle regeneration after minor trauma is a natural process of adaptation to excessive

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stressing and is accomplished by multiple cell types that arrive at the site of injury to begin

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repair [1]. Beyond a certain injury severity threshold this inherent regeneration potential

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proves insufficient [2], leading to fatty degeneration and fibrotic scar tissue formation, which

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compromise muscle function and structural integrity [3]. Such severe muscle damages

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resulting from direct trauma, iatrogenic injuries during surgical procedures, muscle tears,

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ischemia, or myopathies are common [4]. Current treatment options are limited and plagued

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with lack of allogeneic donor tissue or donor site morbidity associated with autograft

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transplantations [5]. Moreover, they often result in unsatisfying functional outcome,

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substantiating the clear medical demand for novel therapeutic approaches [6, 7].

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Strategies aiming to improve muscle regeneration via the local delivery of cells, growth

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factors, or a combination thereof are currently under investigation [8, 9]. Growth factor

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delivery has been shown to be a useful approach to treat ischemic muscle tissues in animal

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models [10], but their effectiveness in the repair of severe injuries is yet to be demonstrated.

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Moreover, the large number of complex biological processes necessary to induce repair of

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injured muscle tissues may require the spatiotemporal delivery of several distinct growth

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factors, which poses a significant challenge and renders this strategy suboptimal [11, 12].

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Since the failure of direct myoblast delivery in earlier clinical trials [13, 14], adult progenitor

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cells, e.g. muscle satellite cells or mesenchymal stromal cells (MSCs), have emerged as

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promising cell types to enhance muscle regeneration [15]. MSCs are highly proliferative

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multipotent cells that are known to modulate injury responses through paracrine signaling

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[16-18]. For instance, MSCs serve as a source of cytokines and proteinases essential to

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angiogenesis and tissue regeneration, such as VEGF, MMPs, IGF-1, HGF, TGF-β, and bFGF

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[19]. The regenerative capability of MSCs has been validated in several animal models of

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muscular dystrophy, myocardial infarction as well as muscle trauma [20-24]. Collectively,

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these studies provide evidence that MSC administration reduces the local inflammation and

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enhances tissue function, not as a result of differentiation into muscle cells and fusion with

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existing myofibers, but apparently due to paracrine effects via the secretion of cytokines and

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other growth factors [25, 26]. Despite potential efficacy, bolus cell transplantation frequently

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suffers from lack of tissue engraftment and very low cell survival at the site of injury,

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compelling researchers to inject higher cell numbers to achieve significant improvements [27,

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28]. Thus, it appears essential for an effective cell therapy to ensure localization of

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transplanted cells at the site of injury, for example by using biomaterial scaffolds.

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A biomaterial driven approach could overcome the problems associated with bolus cell

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injection by ensuring a localized delivery of signaling factors while shielding cells from

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harmful environmental effects of the injury site [29, 30]. Various biomaterials have been

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employed to transplant cells that could undergo myogenic differentiation and fuse with

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existing fibers of the injured tissue [31-34]. The success of such a strategy hinges on a number

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of critical factors including migration of cells from the biomaterial substrate towards the

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injured tissue, successful engraftment and differentiation, and requires a large number of cells

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in case of volumetric muscle loss or severe trauma. However, novel approaches stipulate

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biomaterials that present multiple cues for optimization of cell function [35]. We

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hypothesized that a more effective approach would be to support the endogenous regenerative

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mechanisms of the skeletal muscle by harnessing desirable cytokines and growth factors

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secreted by autologous MSCs and ensuring their local delivery at the site of injury. A suitable

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porous scaffold with drug release capability would provide an appropriate structural and

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chemical environment, in essence mimicking a niche that is conducive to MSC paracrine

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function. It was our aim to show that MSCs need not engraft onto the host tissue and undergo

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differentiation, but could induce muscle regeneration by secreting factors from the niche. For

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this reason, we chose not to use an injectable in-situ forming hydrogel to deliver the cells

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intramuscularly, instead choosing to place the MSC loaded niche adjacent to the injured

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muscle. Moreover, in line with clinical observations, the injured muscle is normally exposed

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and in this case an injectable hydrogel, deliverable in a minimally invasive manner, may not

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offer an advantage over a macroporous scaffold.

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We therefore engineered a porous (70-150 µm) multifunctional alginate cryogel that acts as a

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synthetic niche for MSCs, supports their adhesion and spreading while simultaneously

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boosting their secretory function via local stimulation with IGF-1 and VEGF165. The

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transplantation of this synthetic niche in a clinically relevant crush muscle trauma model

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significantly improved muscle strength, reduced fibrosis, and increased muscle fiber density

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and the number of regenerated fibers. This work shows that without any genetic manipulation,

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MSC’s paracrine factor secretion can be upregulated, the released factors being more potent at

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influencing myoblast behavior and orchestrating muscle regeneration than standalone MSC or

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growth factor delivery.

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2. Materials and Methods

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Animals: 4 month old female Sprague Dawley rats (Charles River laboratories), weighing

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between 200-250 g were used for this study with each group being representative for at least 8

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animals. All animal experiments were carried out in compliance with the policies and

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principles established by the Animal Welfare Act, the NIH Guide for Care and Use of

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Laboratory Animals, and the national animal welfare guidelines. The study was approved by

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the local legal representative (Landesamt für Gesundheit und Soziales, Berlin: Registration

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number: G0119/12)

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MSC isolation and cell culture: Bone marrows were aspirated from rat tibiae, and cultured in

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DMEM containing 10 % FBS and 1 % penicillin/streptomycin. MSC phenotype and

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differentiation potential was confirmed as described earlier [36]. Briefly, cell surface marker

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expression was validated using flow cytometry. MSC populations were positive (≥ 95% of

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total counts) for CD29, CD44, CD105, CD73, CD166, CD90, and RT1A and negative (≥

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98%) for CD45, CD34 and RT1B. Differentiation potential upon stimulation with adipogenic

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or osteogenic media was confirmed via Nile red and Alizarin Red staining, respectively.

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Unless otherwise stated, MSCs from at least four different donors were used in the in vitro

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assay described below.

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The C2C12 cell line was purchased from ATCC (Catalog # ATCC® CRL-1772™).

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Expansion media for C2C12 myoblasts contained 10 % FCS, which was reduced to 5 % to

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induce differentiation.

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MSC conditioned medium: MSCs between passage 3 and 5 were cultured in 6 well plates at a

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density of 3 x 105 cells per well, and allowed to attach. After overnight incubation, cells were

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washed twice with calcium free PBS and 1.5 mL/well of fresh serum free media was added.

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For MSC+GF-CM, the serum free media was supplemented with 5 ng/mL of IGF-1 and

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VEGF165. After a further 24 hour incubation period, the conditioned media was collected and

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centrifuged at 700 rcf for 8 minutes to remove cell debris, and passed through a 0.2 µm sterile

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filter. The conditioned media was then concentrated 20x using Amicon ultracentrifugal filter

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units (10 kDa) and re-diluted in fresh serum free media to its original volume. For

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differentiation studies involving C2C12 myoblasts, supplementary FCS (5% v/v) was added

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to the conditioned media. For growth factor neutralization experiments, conditioned media

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containing IGF-1 and VEGF165 were incubated with anti-human VEGF antibody (R&D

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systems, Clone # 26503, 1:10000) and anti-human IGF antibody (R&D systems, catalog #

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AF-291-NA, 1:500) for 4 hours before use.

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Myoblast proliferation & survival: C2C12 myoblasts were seeded in 48 well plates at a

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seeding density of 3 x 103 per well and allowed to attach. After overnight incubation, media

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was removed and the cells were washed twice with calcium free PBS before adding either (1)

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serum free cell culture media (blank), (2) serum free media containing growth factors at a

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concentration of 5 ng/mL each (+GF), (3) conditioned media from MSCs (MSC-CM), or (4)

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conditioned media from MSCs stimulated with 5 ng/mL growth factors (MSC+GF-CM).

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After 48 hours, media was aspirated and the well plates were frozen at -80 °C. CyQuant® cell

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proliferation assay kit (Invitrogen) was used to determine cell proliferation. Myoblast survival

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was quantified by first seeding 3 x 104 C2C12 cells/well of a 48 well plate. After overnight

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attachment, a scratch was made to mimic injury, washed twice with PBS, and then the

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appropriate media was added. After a further 24 hours, cells were trypsinized after washing

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with PBS. The cell suspension was mixed with trypan blue and viability was determined by

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counting cells in a hemocytometer.

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Myoblast apoptosis rate: Apoptosis was evaluated with Caspase-Glo® 3/7 assay (Promega)

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according to the manufacturer’s instructions. Briefly, myoblasts were seeded on white-walled

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96 well plates with a cell density of 1 x 104/well. After overnight attachment a scratch was

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made and appropriate media were added. After a further 24 hours, a 1:1 mixture of the

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Caspase-Glo® buffer and substrate was added into the wells and luminescence was detected

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using a plate reader. Measurements were normalized to cell number determined by

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CyQuant®. Myoblast survival was quantified by a similar seeding method, but 24 hours after

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application of respective media, cells were trypsinized after washing with PBS. The cell

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suspension was mixed with trypan blue and viability was determined by counting cells in a

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hemocytometer.

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Migration assay: The migration behavior of C2C12 myoblasts in response to different media

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was investigated using a scratch wound healing assay. 1.5 x 105 myoblasts were seeded in

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each well of a 24 well plate and allowed to attach overnight to form a confluent monolayer. A

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200 µL pipette tip was used to create a scratch; wells were washed twice with PBS before

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adding indicated conditioned media. Images were taken every 30 minutes over a period of 24

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hours using an inverted microscope (DMI6000B, Leica, Germany) with a live cell imaging

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system. The images were analyzed using T-scratch [37], and the kinetics of wound closure

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were determined for the different treatment groups.

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Myoblast differentiation (Western Blot): To monitor the influence of MSC’s conditioned

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media on myoblast differentiation behavior, 4 x 105 C2C12 cells were seed in each well of a 6

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well plate. After overnight attachment and the formation of a confluent layer, the appropriate

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conditioned media were added. Four days after induction of differentiation, C2C12 cells were

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washed with PBS, and lysates were generated using SDS lysis buffer. Protein content was

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determined using DC protein assay (Bio-Rad). The Novex NuPAGE® system (Invitrogen)

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was used according to manufacturer’s instructions followed by semi-dry transfer. Primary

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antibodies for Myosin heavy chain (Clone # MF20, R&D Systems, Catalog # MAB4470)

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(1:500), Myogenin (Clone # F5D, Abcam, Catalog # ab1835) (1:500), MyoD (Clone # 5.8A,

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Thermo Scientific, Catalog # MA1-41017) (1:500), and the house keeping gene GAPDH

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(Clone # 6C5, Abcam, Catalog # ab8245) (1:10000) were used with TBST+5% milk. Anti-

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mouse horseradish peroxidase (GE Healthcare) (1:7000) was utilized as the secondary

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antibody. Using ECL substrate (GE Healthcare) pictures were acquired using the G:BOX

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imager (Syngene). Analysis was performed using band intensities normalized to GAPDH and

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quantified by NIH ImageJ software package (http://rsb.info.nih.gov/ij/).

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Myoblast differentiation (Fluorescence microscopy): Cells were seeded at a density of 1 x

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105/well of a 24 well plate. Four days after inducing differentiation in C2C12 myoblast

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cultures, cells were washed with PBS, fixed with 4 % paraformaldehyde, permeabilized with

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0.1 % saponin in 3 % BSA solution, and stained with anti-Myosin heavy chain (1:200), and

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DAPI (1:1000). Cells were imaged with an inverted fluorescence microscope (Leica).

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Alginate processing: Ultrapure alginates were purchased from Novamatrix (Oslo, Norway).

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MVG alginate was used as the high molecular weight component (250 kDa) and LVG

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alginate (50 kDa) was used as the low molecular weight component to prepare gels. Both

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alginate polymers were diluted to 1 % w/v in double-distilled H2O, and 1 % of the sugar

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residues in the polymer chains were oxidized with sodium periodate (Sigma-Aldrich, St

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Louis, MO) by maintaining solutions in the dark for 17 hours at room temperature under

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constant stirring, as previously described [38]. An equimolar amount of ethylene glycol

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(Fischer, Pittsburgh, USA) was added to stop the reaction, and the solution was subsequently

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dialyzed (MWCO 3500 Da, Spectra/Por®) over three days, frozen at -20° C, and lyophilized.

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Both alginates were modified with G4RGDSP peptides (Commonwealth Biotechnology,

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Richmond, USA) at a degree of substitution of 10 (10 peptide molecules per alginate chain)

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using carbodiimide chemistry, as previously described [39]. The modified alginate solutions

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were dialyzed again, mixed with activated charcoal, sterile filtered, lyophilized, and stored at -

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20° C until further use.

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Scaffold Fabrication: Equal amounts of lyophilized MVG and LVG alginates were mixed,

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and reconstituted in high glucose containing DMEM (Sigma-Aldrich, Germany) to obtain a

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4 % w/v polymer solution. Using two syringes coupled with a syringe connector, the alginate

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mixture was ionically cross-linked using aqueous calcium sulfate slurry (40 µL of CaSO4 per

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mL of alginate mixture) by vigorously mixing the contents of the two syringes, to produce

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gels with a final concentration of 2 % w/v. Where appropriate, recombinant human IGF-1 and

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recombinant human VEGF165 (R&D systems, USA) were mixed (at a concentration of 60 µg

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per mL of alginate solution) with the alginate mixture prior to crosslinking. The mixture was

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poured between sterile glass plates separated by 2 mm spacers, and allowed to gel for 30

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minutes. Gels measuring 8 mm x 3 mm x 2 mm were cut out using a punch and frozen at -

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80°C for 24 hours. Thereafter, the frozen gels were lyophilized to induce porosity. For in vivo

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transplantation, 50 µL of a 20 x 106 MSCs/mL cell suspension was pipetted on top of the

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freeze dried porous scaffold and incubated for 20 minutes, before adding excess DMEM.

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Viability of MSCs: The viability of MSCs inside the scaffolds was evaluated over a period of

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14 days using a Live/Dead cell imaging kit (Invitrogen). Briefly, cell seeded scaffolds (1 x 105

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MSCs/scaffold) were rinsed thrice in calcium free PBS before dissociation in a 50 mM EDTA

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solution. The cell suspension was mixed with a mixture of Calcein AM and Ethidium

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homodimer and imaged with a fluorescence microscope after a 15 minute incubation period.

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The number of live (green) and dead (red) cells were counted using NIH ImageJ software

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package.

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MSCs outward migration: To determine outward migration, cell seeded scaffolds (1 x 105

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MSCs/scaffold) were placed in separate wells of a 24 well plate. After every 24 hours, the

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scaffolds were shifted to new wells, and the number of cells that had colonized the wells over

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the previous 24 hours was quantified using the CyQuant® cell proliferation assay kit

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(Invitrogen).

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ELISA: For release kinetics, growth factor containing scaffolds (n=3) were immersed in 1 mL

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of DMEM and incubated for 30 days. At different time points (days 1, 3, 7, 14, 21, and 28),

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supernatants were collected and fresh media was added. The amount of IGF-1 and VEGF165 in

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the supernatants was quantified using Quantikine™ human-specific ELISA kits (R&D

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systems, Germany). Quantitative concentrations of MSC secreted Hgf, bFgf, Igf-1, and Vegf

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in conditioned media were determined using commercially available rat-specific ELISA kits

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(R&D systems) and used according to the manufacturer’s instructions. Cross reactivity with

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human recombinant growth factors was determined (Supplementary information).

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Animal experiments: Animals were randomly allocated into the following treatment groups

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(n=8): (1) empty alginate scaffold, (2) alginate scaffold with incorporated growth factors (3

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µg each of IGF-1 and VEGF165), (3) alginate scaffold seeded with autologous MSCs (50 µL

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of a 20 x 106 MSCs/mL cell suspension), and (4) alginate scaffold with incorporated growth

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factors and seeded with autologous MSCs. Independent from their final group allocation, all

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animals underwent a bone marrow biopsy to harvest autologous MSCs. Bone marrow

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aspirates were taken from both tibiae as described in [40]. Three weeks later, the left soleus

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muscles of the animals were bluntly crushed (monolateral surgery) and the appropriate

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scaffold was immediately transplanted during the same surgery. At day 7, 28, and 56 after

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transplantation, contraction forces of the injured and the contralateral (uninjured) soleus

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muscle were measured. Subsequently, the animals were then sacrificed and the muscles were

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harvested for histological analysis. For the 56 day time point, only groups 1 (Alg.), 3

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(Alg.+MSC), and 4 (Alg.+GF+MSC) were transplanted based on the legal specification of the

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responsible authorities, who only permitted us to monitor the long term effects of treatments

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showing significant difference to the untreated controls at day 28.

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Muscle trauma model and scaffold transplantation: The animals were anesthetized by

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inhalation of isoflurane, administered Rimadyl® subcutaneously, and the lower left limbs

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were shaved. An injury model consisting of standardized blunt crush traumas to the left soleus

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muscle was employed using procedures described earlier [40]. On the day of surgery,

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autologous MSCs were trypsinized and 50 µL of a cell suspension containing 20 x 106

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MSCs/mL was seeded onto each scaffold (total: 1 x 106 MSCs/scaffold). After an initial 20

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minute incubation period, 500 µL of full culture media was added to the well containing the

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cell seeded scaffold. The construct was incubated for a further 4 hours to allow for complete

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infiltration of the cell suspension and to facilitate cell attachment. The scaffold was then

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immediately transplanted into the recipient animal by placing it along the length of the injured

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soleus muscle in a fascia pocket lying in between the soleus and the gastrocnemius muscles.

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No sutures were required to immobilize the scaffold. Thereafter, the superficial muscle and

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skin were closed by suturing, and the wound was cleaned with saline solution.

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Muscle force measurement: After an initial isoflurane narcosis, rats were anesthetized by

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intraperitoneal injection of a mixture of xylazine and ketamine in saline before proceeding

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with the muscle force measurement and muscle harvesting. Fast twitch and slow twitch

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(tetanic) forces of the treated and contralateral uninjured muscles were measured as described

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earlier [28]. Briefly, after anesthetizing the animals, the sciatic nerve and the soleus muscle

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were exposed whilst care was taken to protect the neuromuscular junctions and the

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neurovascular bundle. The Achilles tendon was severed and the lower extremity was

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connected to the muscle force measuring device using a silk suture. Subsequently, the sciatic

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nerve was electrically stimulated and the contraction forces were recorded using a force

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transducer. After the muscle force measurements, the animals were sacrificed.

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Histology: Soleus muscles embedded in Tissue-Tek® O.C.T.™ compound (Sakura), were

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divided into 9 segments along the length of the muscle, and 10 consecutive slices (7 µm

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sections) from each segment were sectioned and fixed onto glass slides (Marienfeld). Sections

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undergoing H&E and Picrosirius red staining were post-fixed with 4 % formaldehyde. To

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identify collagenous scar tissue (Sirius red), the sections were stained with Direct Red 80

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(Fluka, Germany) by incubation in Sirius red solution (1% w/v in saturated picric acid) for 60

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minutes. Differentiation was reached with two washes in diluted acetic acid. Dehydration was

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carried out by immersion in a graded series of ethanol before being incubated in xylol (T. J.

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Baker) twice for 5 minutes. The amount of fibrosis in muscle sections was quantified using a

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custom built macro in NIH ImageJ software package. The fibrotic area in the treated muscle

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was expressed as a percentage of the total section area, and normalized to the percentage of

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connective tissue in the uninjured control. Muscle sections were stained with Hematoxylin

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and Eosin (Merck, Germany) for visualization of myofiber nuclei and cytoplasm. For nuclei

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staining, sections were incubated in Hematoxylin for 7 minutes then washed twice in distilled

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water. Differentiation was achieved by shortly placing the sections in 0.25 % HCl-ethanol

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solution (Merck, Germany) followed by washing under tap water for 10 minutes. For

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cytoplasm staining, sections were placed in 2 % Eosin (CHROMA) for up to 3 seconds and

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dehydrated in graded series of ethanol and finally incubated in xylol twice, each time for a

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duration of 5 minutes. The number of fibers with centrally located nuclei was counted

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manually and expressed as a percentage of the total section area. Muscle fibers expressing

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slow myosin were detected with monoclonal anti-myosin (skeletal, slow) antibody (1:10000)

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(Sigma-Aldrich, clone # NOQ7.5.4D, catalog # M8421). Blood vessels were detected with

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CD31 monoclonal antibody (1:50) (abcam, clone # TLD-3A12, catalog # ab64543). For both

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stainings, air dried sections were fixed in cold acetone for 20 minutes and washed in 10 %

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PBS twice for 5 minutes each. Sections were blocked in 2 % horse serum (Biozol) diluted in

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2 % BSA-PBS for 30 minutes at room temperature. The primary antibodies were then applied

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with overnight incubation at 4 °C. After washing twice with PBS, the secondary antibody

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(Biotinylated Horse Anti-Mouse IgG Antibody, rat adsorbed, Biozol) was applied (1:50) for

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30 minutes. The sections were washed twice with PBS and the avidin-biotin complex was

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applied for 50 minutes at room temperature (AP-Standard kit AK 5000; Vector). After

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washing, sections were then incubated with Alkaline Phosphatase substrate (Red Alkaline

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Substrate Kit I, SK-5100, Vector) for up to 8 minutes. Finally, sections were counter-stained

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following the Mayers Haemalum method. Slow fibers were quantified using a custom built

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macro in NIH ImageJ software package. For quantification, positively stained blood vessels

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were manually counted from regions of interest generated randomly from the entire muscle

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section. For all stained sections, images were captured using a light microscope (Leica

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Microsystems, Germany) equipped with a digital camera (AxioCam MRc, Carl Zeiss

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MicroImaging, Germany).

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Statistical analysis: The IBM® SPSS® 22.0 software package (IBM Corp., Chicago, IL,

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USA) was used for statistical evaluation. Results are presented as mean ± standard error of the

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mean (s.e.m). Unless otherwise stated, all in vivo samples are representative of at least n = 8

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animals (per group & time-point, range: 8-10) and all in vitro results are representative of at

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least four biological replicates analyzed in two independent experiments. Sample size was

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selected based on previous experience [28, 40] to minimize the number of animals needed

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while obtaining a valid statistically significant result (in accordance with the 3R principles).

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The Shapiro-Wilk test was used to test normality and the Levene test was used to assess the

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homogeneity of variances of the data for the indicated groups. Independent and non-normal

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distributed data were analyzed using the Mann-Whitney U test (two groups) or the Kruskal-

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Wallis one-way analysis of variance (>two groups). If normality could be assumed, unpaired

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Student’s t-test (two groups) or the corresponding one-way analysis of variance (>two groups)

284

was chosen to determine statistical significance. P-values were adjusted using Bonferroni’s

285

(equal variances) or Dunnett T3 (unequal variances) p-value adjustment multiple comparison

286

procedure. Dependent sample data were analyzed using Wilcoxon signed rank and paired

287

Student’s t-test (two samples), respectively, or Friedman’s two-way ANOVA by ranks (> two

288

groups). All tests were analyzed two-sided and P<0.05 was regarded as significant. Detailed

289

information about error bars, sample sizes per group, and statistical analysis are included in

290

all figure legends. The investigators were not blinded to group allocation during the

291

experiments because the described treatments displayed obvious differences.

292

3. Results

293

3.1. Effect of MSC derived paracrine factors on myoblast function

294

The potential effects of bioactive factors derived from rat MSCs on muscle progenitor cells

295

was examined by exposing skeletal myoblasts (C2C12) to conditioned media generated from

296

unstimulated (MSC-CM), and recombinant growth factor (IGF-1 & VEGF165) stimulated

297

MSCs (MSC+GF-CM). Assessment of the growth dynamics of myoblasts via the number of

298

population doublings (PD) showed higher proliferation rates for C2C12 cells grown in MSC-

299

CM or MSC+GF-CM compared to corresponding cultures in serum free media without

300

(blank) or with recombinant growth factors (+GF) (Figure 1A). (PD relative to blank:

301

PD+GF=0.97±0.03,

302

PDMSC+GF-CM=1.49±0.05, Pvs.blank<0.001, Pvs.+GF<0.001). The anti-apoptotic benefits of MSC

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Pvs.blank=1.000;

PDMSC-CM=1.43±0.03,

Pvs.blank<0.001,

Pvs.+GF<0.001;

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derived paracrine factors were validated under serum starvation. This analysis revealed

304

significantly improved survival of cells cultured in media +GF (1.7-fold, P<0.001), MSC-CM

305

(2.2-fold, P<0.001), and MSC+GF-CM (2.3-fold, P<0.001) compared to their counterparts

306

grown in blank media. We tested whether improved survival correlated with lower apoptosis

307

rates by determining caspase-3/7 activity as a surrogate marker for apoptotic cell death.[30]

308

Caspase-3/7 activity of C2C12 cells cultured in MSC+GF-CM was 39% (P<0.001) lower than

309

in the parallel cultures with blank media with or w/o GF. Notably, the anti-apoptotic effect of

310

MSC+GF-CM was also significantly higher compared to CM of untreated MSCs (P=0.005),

311

whereas no significant differences in apoptosis rates were observed between MSC-CM and

312

both control groups.

313

For further investigation of the paracrine effects of MSCs on myoblast function, we compared

314

the migration and differentiation potential of C2C12 cells depending on the culture

315

conditions. Employing a scratch wound healing assay to mimic injury, it was found that the

316

supplementation of human GF as well as MSC-CM significantly improved myoblast cell

317

migration (Pvs.blank<0.001; Pvs.+GF<0.001), and that this effect was further enhanced when

318

MSC+GF-CM was applied (Pvs.blank<0.001; Pvs.+GF<0.001; Pvs.CM<0.034) (Figure 1B and C).

319

In contrast, MSC-CM and MSC+GF-CM significantly inhibited myoblast differentiation

320

compared to control media with and without human GF. Cells cultivated in MSC-CM or

321

MSC+GF-CM showed a significantly lower expression of myosin heavy chain (Myh2) and

322

myogenin compared to corresponding control cultures (Figure 1D and E). Accordingly,

323

qualitative analysis by indirect immunofluorescence showed that C2C12 cells underwent

324

terminal differentiation (fusion into multinucleated myotubes and accumulation of Myh2) in

325

the presence of differentiation media independent from GF supplementation, while

326

differentiation was significantly reduced in the presence of MSC-CM or MSC+GF-CM,

327

respectively (Figure 1G).

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These results indicate that paracrine factors secreted from unstimulated and stimulated MSCs

329

promote the proliferation, survival, and migration of C2C12 myoblasts, while preventing their

330

cell cycle withdrawal and subsequent differentiation into myotubes. Previous studies have

331

suggested that promotion of myoblast proliferation and migration, and delaying their

332

differentiation represent favorable conditions for in vivo muscle repair [41, 42].

333

3.2. Modulation of MSC secretion pattern and therapeutic effects by stimulation with

334

recombinant IGF-1 and VEGF165

335

Since GF-stimulation of MSCs enhanced their paracrine actions on myoblasts, we wondered

336

whether this reflected an additive or synergistic effect. Hence, we examined whether

337

depletion of the human GFs from the MSC+GF-CM alters its effects on myoblast

338

proliferation, survival, and migration (Figure 2A, B and C). IGF-1 and VEGF165 were

339

depleted by using human-specific neutralization antibodies and this procedure efficiently

340

reduced levels of the human cytokines in the conditioned media (Figure S1). In the presence

341

of the neutralization antibodies, the positive effects of blank media supplemented with human

342

GF on C2C12 viability and migration were completely abolished. In contrast, the enhanced

343

proliferation, survival, and migration in response to MSC+GF-CM remained significant,

344

confirming that recombinant growth factors do not have an additive effect, but in fact

345

contribute primarily by stimulating MSCs potentially by modulating their paracrine secretion

346

pattern. This was verified by using rat-specific immunosorbent assays (ELISA) to detect

347

cytokines known to play a vital role in muscle regeneration. Conditioned media from GF-

348

stimulated MSCs was found to contain increased levels of basic fibroblast growth factor

349

[bFgf] (1.4-fold, P=0.045), Vegf (1.5-fold, P=0.001), and hepatocyte growth factor [Hgf]

350

(2.4-fold, P=0.026) compared to MSC-CM, while Igf-1 levels were not altered (P=0.277)

351

(Figure 2D). Vegf, bFgf, Hgf, and Igf represent factors originating from rat MSCs, as opposed

352

to recombinant human IGF-1 and VEGF165. Apart from these specific cytokines, the total

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amount of secreted protein was also found to be clearly increased in CM of GF-stimulated

354

MSCs (Pvs.CM=0.006). This might indicate the amplified secretion of other cytokines and

355

growth factors in the conditioned media of stimulated MSCs. Taken together, growth factor

356

stimulation enhances MSC’s paracrine activities and the resulting release of bioactive factors

357

may exert a strong effect on the cells of the injured muscle tissue.

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3.3. Porous alginate cryogel supports MSC attachment and spreading

360

Subsequently, we engineered a multifunctional alginate scaffold that could be utilized as an

361

artificial niche for MSCs at the site of injury (Figure 3A). The niche should allow adhesion

362

and spreading of transplanted MSCs (Figure 3C), protect them during transplantation in an

363

initially inflammatory environment, and allow to boost their paracrine signaling by local

364

release of incorporated human growth factors (IGF-1 & VEGF165). The fabrication of alginate

365

scaffolds consisted of several steps including oxidation of alginate and RGD peptide

366

conjugation. The resulting biomaterial was combined with IGF-1 and VEGF165 before being

367

ionically crosslinked using a calcium source, and lyophilized to produce macroporous

368

scaffolds (Figure 3B). Scaffolds provided a sustained release of incorporated growth factors

369

over a period of 30 days after an initial burst release (Figure 3D). Independent from GF

370

incorporation, MSCs were found to adhere to the walls of the RGD-peptide conjugated

371

alginate scaffold and maintain high viability for at least 7 days (Figure 3E). Although the

372

biomaterial exhibited a porous microstructure (pore size 70-150 µm) only a small percentage

373

(3-6 %) of the seeded cells migrated out of the scaffold and GF incorporation even further

374

reduced this outward migration compared to the blank biomaterial (P<0.001 at all time-points

375

investigated; Figure 3F). Such a scaffold would allow us to test the hypothesis that

376

transplanted MSCs are not required to physically engraft onto the injured tissue to achieve

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regeneration, but could orchestrate the regeneration process by secreting bioactive factors in

378

the injury environment.

379

3.4. Transplanted synthetic niche enhances contraction forces of injured muscles

381

For in vivo investigations, an established, clinically relevant rat model of severely crushed

382

soleus muscle tissue was used [24]. Animals were randomly assigned to different groups and

383

were treated with either (1) blank scaffold (Alg.), (2) scaffold containing growth factors

384

(Alg.+GF), (3) scaffold seeded with autologous MSCs (Alg.+MSC), or (4) scaffold

385

containing both MSCs and growth factors (Alg.+GF+MSC). Quantitative measurements of

386

fast twitch and tetanic muscle forces were performed after 7, 28, and 56 days post injury using

387

a custom made set up that stimulated the sciatic nerve and recorded muscle contractions via a

388

force transducer, as described earlier [28]. Two additional regimes were tested to determine

389

their functional benefit: (a) bolus intramuscular injection of growth factors (inj. GF) and (b)

390

bolus intramuscular injection of MSCs and growth factors (inj. MSCs+GF); data shown in

391

supplementary information (Figure S2). All muscle forces were normalized to the uninjured

392

contralateral measurements.

393

At day seven, we found a comparable reduction of the fast twitch (~57±2%) and tetanic

394

(~46±2%) muscle force between all injured muscles (Figure 4A). At day 28, the fast twitch

395

muscle force progressively increased from Alg. (58±4%), Alg.+GF (64±3%), Alg.+MSC

396

(74±4%, PAlg.=0.024), to the highest value detected in the Alg.+GF+MSC group (83±3%,

397

PAlg.<0.001, PAlg.+GF=0.003). We observed a similar trend toward increased tetanic muscle

398

force, however, this tendency only reached statistical significance between Alg.+GF+MSC

399

and the alginate control (P=0.018) (Figure 4B). In order to investigate the long lasting effects

400

of the most promising groups (Alg.+MSC & Alg.+GF+MSC) relative to control (Alg.), we

401

additionally determined their muscle forces after 56 days post injury. Fast twitch and tetanic

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forces in the Alg.+GF+MSC group were significantly higher compared to the Alg. control,

403

whereas the tetanic contraction were significantly elevated in both the Alg.+MSC and

404

Alg.+GF+MSC groups in relation to the Alg. control.

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3.5. Effect of transplanted niche on regeneration of muscle fibers

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To determine which structural aspects are affected by the paracrine signaling and contribute to

408

functional benefit, the muscle fiber density and number of regenerated muscle fibers were

409

quantified in multiple histological cross sections distributed over the entirety of the muscle

410

tissues.

411

In line with the data from the muscle force measurement, no difference in the total number of

412

fibers per section area (muscle fiber density) was observed seven days after injury (Figure 5

413

A; Figure S4). However, the fiber densities in the Alg. (-44%, P<0.001) and the Alg.+GF

414

( 26%, P=0.016) groups were significantly decreased at day 28 compared to day seven,

415

whereas no significant change was observed in the Alg.+MSC (-14%) and Alg.+GF+MSC

416

(+17%) group. Accordingly, treatment of muscles with Alg.+MSC or Alg.+GF+MSC, led to a

417

1.7-fold (P<0.001) and 2-fold (P<0.001) higher fiber density, respectively, compared to the

418

blank alginate group at day 28. Moreover, the fiber density in the Alg.+GF+MSC group was

419

also significantly higher compared to Alg.+GF group (1.5-fold, P=0.001). At day 56, the

420

combination of Alg.+GF+MSC showed a significantly higher muscle fiber density compared

421

to both the Alg. and the Alg.+MSC groups. These results suggest that transplantation of the

422

MSC and GF loaded synthetic niche most likely enhances the survival of the muscle

423

progenitor cells at early phases of the healing process, and may promote fusion and muscle

424

fiber formation at later time points [43].

425

To validate this hypothesis, we investigated the number of regenerated fibers identified by

426

centrally located nuclei within the muscle cross sections. Indeed, the number of regenerated

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fibers per section area was quantitatively higher in muscles treated with the Alg. and Alg.+GF

428

groups compared to the Alg.+MSC and Alg.+GF+MSC groups at day seven (Figure 5B). At

429

day 28 however, we observed a trend towards increased number of regenerated fibers in

430

muscles treated with Alg.+MSC and Alg.+GF+MSC combinations, but this trend did not

431

reach statistical significance. However, the muscles treated with the combination of

432

Alg.+GF+MSC showed a significantly increased number of regenerated fibers compared to

433

the two other groups at day 56 (Figure 5C).

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434

3.6. Stimulation of angiogenesis and remodeling of scar tissue

436

Next to muscle tissue changes during regeneration, we quantified the amount of fibrotic scar

437

tissue as well as the blood vessel density in the treated muscle tissues. The amount of fibrotic

438

scar tissue was quantitatively lower in muscles treated with the Alg.+GF+MSC combination

439

compared with both control groups (Alg. or Alg.+GF) at all time points investigated (Figure

440

6A and B). In particular, at day seven after injury the amount of fibrotic scar tissue

441

progressively decreased from Alg.+GF (23.8±1.4%), Alg. (20.3±1.0%), Alg.+MSC

442

(18.7±1.7%), to the lowest value detected in the Alg.+GF+MSC group (14.8±1.3%, =0.014,

443

PAlg.+GF<0.001). At day 28, muscles treated with Alg.+GF+MSC (14.3±1.3%) showed

444

significantly lower amounts of fibrotic scar tissue than the Alg. group (22.4±1.6%, P=0.003)

445

as well as the Alg.+GF group (22.3±1.6%, P=0.003). A similar tendency could be observed

446

between the Alg.+MSC (17.5±2.4%) and the control groups, but did not reach statistical

447

significance. The positive effect of the Alg.+GF+MSCs therapy seems to be long-lasting,

448

since we also observed significantly lower fibrosis in that group (11.7±1.3%) compared to

449

Alg.+MSC group (19.5±1.5%, PAlg.+MSC+GF=0.001) as well as the Alg. group (25.4±1.5%,

450

PAlg.+MSC+GF>0.001, PAlg.+MSC=0.027) at 56 days after injury (Figure S5).

451

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To test whether improved muscle force induced by the combined administration of

453

Alg.+GF+MSC also corresponded to increased vascularization, the total number of vessels

454

per section area was quantified using CD31 staining (Figure 6C and D). We found a

455

significantly higher number of vessels in muscles treated with MSCs or GF alone as well as

456

with the combination of both, compared to the blank alginate control at all time points

457

investigated (Figure S6).

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Interestingly, we noted the appearance of new muscle fibers and blood vessels in regions

460

predominantly composed of scar tissue even at day 7 after injury (Figure 7A and B). In line

461

with the reduced amount of scar tissue, vessels and muscle fiber formation were markedly

462

higher in animals that received the Alg.+GF+MSC or the Alg.+MSC treatment (Figure S7 and

463

S8). At 56 days after transplantation, the scar tissue had been largely mitigated as the greater

464

tissue area was remodeled by mature, regenerated myofibers in the Alg.+GF+MSC group.

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4. Discussion

467

The data presented here illustrates the superior effect of an engineered niche that modulates

468

MSC’s paracrine secretion, leading to reduced scar formation, increased angiogenesis, faster

469

reconstitution of muscle structure and improved functional outcome after severe muscle

470

trauma.

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MSC-based bolus transplantation strategies represent a viable approach to improve muscle

473

regeneration, but an unfortunate reality of this approach is the dramatically low percentage of

474

transplanted cells that survive the hostile injury environment and an even lower percentage of

475

cells that eventually differentiate into the desired lineage [44]. Despite these drawbacks,

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studies report beneficial effects of direct MSC injection, leading to the speculation that the

477

observed outcomes are due to MSC’s trophic, paracrine or immunomodulatory functions [45,

478

46]. Exploring the ability of MSCs delivered in macroporous scaffolds to reinforce

479

endogenous muscle regeneration by influencing the activity of muscle progenitor cells, our

480

study focuses on MSC paracrine effects independent of their physical engraftment to host

481

tissue, myogenic differentiation, or immune modulation.

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We identified key modulators of MSC paracrine signaling that indirectly facilitate control

484

over myoblast behavior and function in vitro, and transferred this concept to an in vivo

485

validation experiment in a multiple crush trauma model. In contrast to other frequently used in

486

vivo models, including lacerations, needle punctures or toxin-induced injuries, this model is

487

representative of muscle injuries frequently encountered in the clinic [47]. Similar to human

488

data [48], this preclinical model shows not only a loss in muscle function, but also structural

489

degradation of muscle fibers, formation of scar tissue and disruption of the inherent vascular

490

network. In this context, the outcomes of our study represent a significant advancement in the

491

field of muscle regeneration and has the potential for translation into the clinic.

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Animals in all treatment groups exhibited a 50-60% loss in fast twitch and 40-50% loss in

494

slow twitch tetanic muscle forces 7 days post-trauma relative to the contralateral side. The

495

relatively poor functional outcome of the empty alginate group indicates the inability of the

496

natural healing processes to meet the demands created by severe trauma. In stark contrast,

497

exogenous delivery of MSCs recorded significantly higher muscle contraction forces in the

498

long term. Most importantly, co-delivery of MSCs and growth factors resulted in the fastest

499

and largest recovery of muscle forces. These improved functional outcomes were likely a

500

consequence of significant structural remodeling of the treated tissues. In contrast to earlier

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studies that employed an ischemic muscle injury model [10], the GF delivery alone did not

502

lead to significant functional or structural improvements. The positive influence of growth

503

factors on tissue regeneration is frequently described in animal models that involve ischemic

504

injury. In those cases, growth factors like VEGF, Ang1, and FGF play a major role in

505

restoring blood vasculature in the injured tissue [49]. However in our injury model, apart from

506

disruption of blood supply, other issues include the onset of fibrosis, loss of cells, and

507

necrosis of muscle fibers. Despite being delivered in large quantities, it seems that VEGF and

508

IGF were unable to do much in the injured tissue except inducing angiogenesis (Figure 6C,

509

D). On the other hand, the MSCs, being a potent source of multiple cytokines, could

510

orchestrate the numerous biological processes necessary to reverse the effects of severe

511

injury, in addition to angiogenesis.

512

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In vitro, we found that physiologically relevant indicators of an active regeneration process,

514

such as myoblast proliferation, migration, and survival in harsh conditions, were enhanced

515

after culture in MSC-CM and MSC+GF-CM, in contrast to standalone growth factor

516

supplementation that registered modest improvements. By employing neutralization

517

antibodies, we confirmed that recombinant growth factors do not have an additive effect, but

518

in fact contribute primarily by stimulating MSCs, upregulating their paracrine secretion

519

pattern, and consequently improving myoblast function. Correspondingly, analysis of

520

supernatants revealed that GF-stimulation significantly increased the overall protein content

521

secreted by the MSCs and particularly upregulated the secretion of Hgf, Vegf, and bFgf in

522

comparison to MSC-CM. The growth factor release kinetics outcome suggested that most of

523

the incorporated VEGF165 is released from the cryogel within the first 10 days, whereas the

524

release of IGF-1 persists for 30 days. Together with in vitro studies which revealed that MSCs

525

demonstrate enhanced factor secretion after being stimulated with growth factors for only 24

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526

hours, this may indicate that long-term stimulation of MSCs (i.e. more than 10 days) is not

527

necessary to achieve the desired regeneration observed in the Alg.+GF+MSC group.

528

MSCs secrete many factors, which enhance cellular functions including cell division and

530

motility, and provide protection from apoptosis under challenging conditions [25, 50]. For

531

instance HGF, bFGF, GM-CSF, IGF-1, and VEGF have not only been implicated in rendering

532

anti-apoptotic effects [45, 51], but also in the regulation of myoblast cell cycle withdrawal

533

[42, 52-55]. Our results from the Caspase 3/7 activity measurement as well as the

534

differentiation assays are in agreement with the detection of some of these factors in MSC-

535

CM and MSC+GF-CM. In the context of major muscle trauma which results in massive cell

536

loss, our results suggest that MSC-derived factors would likely contribute towards a)

537

preventing further cell loss in the compromised tissue, b) promoting proliferation to increase

538

cell numbers, c) supporting the migration of these proliferating cells to the site of injury, and

539

d) inhibiting the formation of prematurely differentiated muscle fibers. Accordingly, it has

540

been shown that the modulation of the injury environment via soluble factors or anti-fibrotic

541

agents (e.g. TGF-β antagonists) could improve in vivo muscle regeneration by extending

542

myoblast proliferation, increasing their migration and concurrently delaying their

543

differentiation [15, 41, 56].

544

This correlates with our in vivo observation that animals that received Alg.+MSC+GF

545

treatment exhibited the lowest ratio of regenerated fibers at the earliest time point, but

546

recorded a progressive increase thereafter with a significant difference to all other groups at

547

day 56. In accordance with the functional outcome, this treatment also resulted in the highest

548

fiber density of all groups at days 28 and 56. In contrast, animals that received Alg. or

549

Alg.+GF showed quantitatively the highest number of regenerated fibers at the earliest time

550

point, which decreased thereafter. Although no differences in myofiber density were evident

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between all groups at the early time point, we noted that the myofiber density in the Alg. and

552

Alg.+GF group significantly decreased between days 7 and 28. This indicates that the

553

endogenous healing process leads to formation of new fibers, likely due to the action of

554

resident satellite cells, but these fibers fail to reach maturity and become necrotic over time.

555

To the best of our knowledge, there exist no studies investigating the effect of conditioned

556

media from stromal cells on the function of primary satellite cells. This is likely because long-

557

term in vitro culture of satellite cells remains a challenge. Thus, the regenerated fibers in

558

Alg.+GF+MSC group most likely matured over time and participated in the skeletal muscle’s

559

contractile machinery. Additionally, this treatment resulted in a faster restoration of the

560

predominantly slow fiber muscle phenotype, which is also the composition of the native

561

soleus muscle (Figure S3).

562

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Re-establishment of a vascular network is an essential criterion for successful muscle healing

564

[57], as cells are normally unable to survive if a nutrient source is located more than 150 µm

565

away. Analysis of muscle sections stained with CD31 revealed a significantly lower amount

566

of blood capillaries in the Alg. group relative to all other treatments. This suggests that

567

improved vessel formation might not be the leading cause for the improved muscle function

568

after MSCs transplantation. However, the absence of a functioning vasculature may also

569

contribute to lower fiber densities and consequently lower muscle contraction forces. Perhaps

570

more importantly, we also observed the presence of blood vessels in regions predominantly

571

composed of scar tissue. Even at the earliest time point, the number of vessels within the scar

572

tissue was notably higher in animals treated with Alg.+MSC+GF or Alg.+MSC. Scar tissue is

573

thought to establish a permanent presence in injured tissues, but the existence of a vascular

574

network within fibrotic regions pointed towards imminent remodeling. We observed that

575

while the area of scar tissue was significantly reduced over time in Alg.+MSC+GF treated

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animals, there was a gradual increase in the Alg. group. This could imply that a) the natural

577

healing process is unable to curb or mitigate the progression of scar tissue formation, and b)

578

the availability of soluble factors secreted from MSCs even at very early time points induces a

579

dramatic remodeling of the scar tissue in the long-term. For example, the secretion of HGF

580

was significantly enhanced in MSCs upon their stimulation with growth factors; this has been

581

previously described to play an important role in scar tissue remodeling [58, 59]. To the best

582

of our knowledge, there are no existing reports on the effect of MSC derived paracrine factors

583

on the function of ECM secreting fibroblasts or native fibro/adipogenic progenitor cells

584

(FAPs), despite the fact that MSCs secrete a number cytokines with anti-scarring and matrix

585

remodeling properties. It is probable that in addition to influencing myoblast function, MSCs

586

also guide the behavior of fibroblasts and modulate their collagen deposition.

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587

5. Conclusion

589

Collectively, our results suggest that MSC mediated paracrine signaling can support

590

endogenous muscle healing processes in a robust manner, and that the functional and

591

structural outcomes of regeneration can be further improved by enhancing the secretion

592

pattern of MSCs via local growth factor stimulation. The synthetic niche approach described

593

in this study is an important example of enabling in situ regeneration of tissues without

594

physical engraftment of transplanted cells, and will likely be an approach that is applicable in

595

the treatment of a number of compromised tissue functions.

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Supporting Information

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Supporting Information accompanies this paper.

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Acknowledgements

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We would like to acknowledge Gabriela Korus and Mario Thiele for providing technical

602

assistance with histology, and ImageJ macros, respectively. Funding: This work was

603

supported by grants from the BCRT and BSRT through funding by the German Federal

604

Ministry of Education and Research (BMBF), by a grant (GE2512/1-2) from the German

605

Research Foundation (DFG) to SG, and by the award of the Einstein fellowship to DJM. THQ

606

wishes to acknowledge funding from the Friede Springer Foundation and the BSRT. The

607

funders had no role in study design, data collection and analysis, decision to publish, or

608

preparation of the manuscript.

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Figure 1. Paracrine factors in conditioned media influence myoblast function in vitro. (A) Conditioned media from unstimulated and stimulated MSCs enhance myoblast proliferation (CyQuant®), survival (Trypan Blue), and prevent their apoptosis (Caspase-Glo® 3/7) [n=10,*P<0.05, ANOVA with Bonferroni correction]. (B) Assessment of migration behavior of C2C12 cells in a scratch wound healing assay revealed significantly faster migration in response to MSC+GF-CM. Graphs illustrate the kinetics of scratch repopulation over time (left), and the corresponding single point evaluation at 20h (right). [n=8,*P<0.05, ANOVA with Dunnett correction]. (C) Representative images of the migration assay taken using an inverted live cell-imaging microscope 20h after scratching the cell monolayer. (D-E) Myoblast differentiation quantified using Western blot analysis of Myosin Heavy Chain (Myh2), MyoD, and Myogenin. Equal protein loading was confirmed by detecting GAPDH. The results were quantified by densitometry and are shown in the corresponding diagram. [n=4, *P<0.05, ANOVA with Dunnett correction]. (F) Indirect immunofluorescence of Myh2 (green) and DAPI (blue) showed that C2C12 cells cultivated in blank and blank+GF groups highly expressed Myh2 and formed multinucleated myotubes, while fusion was inhibited in the presence of MSC-CM or MSC+GF-CM. Error bars in all panels represent standard error of the mean.

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Figure 2. Growth factors indirectly influence myoblast function by altering secretion pattern of MSCs. Depletion of IGF-1 and VEGF165 did not alter (A) proliferation, (B) survival, or (C) migration of C2C12 cells cultured in MSC-CM or MSC+GF-CM, whereas positive effects of blank media with GF (+GF) receded. (D) MSCs upregulated their protein and cytokine secretion in response to GF stimulation. MSC+GF-CM contained higher levels of Hgf, bFgf, and Vegf. (For [A - D] n=5, *P<0.05, [A & B] ANOVA with Bonferroni correction, [C] ANOVA with Dunnett correction, [D] paired two-sample t-test). Error bars in all panels represent standard error of the mean.

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Figure 3. An engineered synthetic niche provides MSCs with a structural and chemical environment that is optimal for paracrine secretion. (A) The schematic illustrates the strategy employed in this study. A porous scaffold containing MSCs and growth factors was placed adjacent to the injured muscle. MSCs in the porous niche are stimulated by the sustained release of growth factors, resulting in the secretion of paracrine factors that promote muscle regeneration. (B) Representative SEM image showing the macroporous structure of the alginate scaffold. (C) Representative fluorescent image of rat bone marrow derived MSCs (stained with CellTracker™ and DAPI) 24 hours after seeding on the scaffold. (D) Release kinetics of IGF-1 and VEGF165 from the alginate scaffolds over a period of 30 days monitored using ELISA. (E) Independent from GF incorporation, MSCs remained highly viable for at least 7 days in vitro. (F) Emigration of cells from the scaffolds was quantified using CyQuant® assay. The results indicated that a very low percentage of cells inside the scaffold migrated out over a period of 7 days. (For [D - F] n=5, [E & F] ANOVA with Bonferroni correction). Error bars in all panels represent standard error of the mean.

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Figure 4. Recovery of contraction forces in treated muscles. Functional regeneration was assessed by measuring (A) fast twitch, and (B) tetanic contraction forces at three time points post injury. Forces were normalized to uninjured contralateral muscles, and the graphs show the relative fast twitch and tetanic forces. Animals that received the Alg.+GF+MSC transplantation exhibited the largest recovery of contraction forces. [n ≥ 8, *p<0.05, ANOVA with Bonferroni correction]. Error bars in all panels represent standard error of the mean.

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Figure 5. Structural repair accomplished via new myofiber formation. (A) Muscle fiber densities were calculated by normalizing the total number of fibers to section area, and revealed higher mature fibers in animals treated with Alg.+GF+MSC in the long term. (B) The number of regenerated fibers (identified by their central nuclei) corroborated the fiber density measurements in the different groups over time. The results indicate a continuous remodeling of the muscle tissue structure. (C) Representative images of myofibers (crosssectional view) over time. Scar tissue is visibly present in the Alginate group even 56 days after injury. This is notable reduced (Alginate+MSC) or absent (Alg.+GF+MSC) in the other treatment groups. The growth factor treated group (Alg.+GF) was not proceeded to 56 days, and is shown in Figure S9 and S10. (n ≥ 8, *p<0.05, [A] ANOVA with Bonferroni correction, [B] Kruskal-Wallis Test with adjustment for multiple comparison). Error bars in all panels represent standard error of the mean.

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Figure 6. Blood vessels that infiltrate fibrotic areas and facilitate myofiber formation expedite scar tissue remodeling. (A) Fibrotic scar tissue area was quantified over time and revealed dramatic remodeling in animals treated with Alg.+GF+MSC. (B) Representative postprocessed histological images of muscle sections showing scar tissue regions (dark red). Sections from the Alg.+GF group are shown in Figure S9. (C) Vascularization of injured tissues was analyzed with CD31 staining that revealed significantly low number of blood vessels in the control (Alg.) group at all time points. (D) Representative images of vascularization in injured tissues after different treatments (blood vessels stained dark red). Sections from the Alg.+GF group are shown in Figure S9. (For [A, C] n ≥ 8, *p<0.05, [A] ANOVA with Dunett T3 correction, [C] ANOVA with Bonferroni correction). Error bars in all panels represent standard error of the mean. Figure 7. Presence of regenerated muscle fibers and blood vessels in the scar regions of the injured tissues (pale pink = scar tissue, bright pink = muscle fibers). (A) Representative images of scar tissue remodeling by new myofibers indicate structural benefits of MSC derived paracrine factors which are notably absent in the control (Alg.) group even after 56 days. (f) We propose this remodeling of scar tissue was facilitated by infiltration of blood vessels into the fibrotic regions (blood vessels stained dark red).

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