0.5 to 1.9 GPa, ultimate tensile strength from 50 to 120 MPa and strain at break between 5 and 20% [100] [101]. Scaffolds must meet as much as possible these properties to resist solicitations and promote cell differentiation [102]. Even though tendons and ligaments are sometimes indiscriminately named in papers given their similarities, they do exhibit differences when it comes to mechanical properties. Ligaments are usually not as stiff (modulus from 5 to 450 MPa) and strong (UTS between 1 and 50 MPa) as tendons and are more elastic (strain at break from 10 to 40 %) [101]. Another specificity that the scaffolds must mimic is the uniaxial structure of tendons, aligned along the direction of load. To obtain neo-tendons by promoting cell alignment and differentiation, scaffolds should mimic this anisotropy. The influence of scaffold anisotropy has been widely investigated, by using microgrooves in the micrometer scale for instance [103][104]. Scaffold alignment may be important down to 100 nm, based on the work of Van Delft and co-workers [105].

Scaffolds for reconstruction must also comply with general specifications for regenerative scaffolds. Biocompatibility is an obvious specification but its definition is not straightforward and has evolved with time. Now, it goes further than the old definition, i.e. the absence of immune rejection or toxicity, called “biotolerable” by Mertz [106], adding “the ability of a device material to perform with an appropriate host response in a specific situation” [107]. Scaffolds should not be inert to the body but bioactive, promoting cell adherence and proliferation. Scaffold porosity is here paramount to ensure exchanges and cell penetration. Besides, the scaffold must be regenerative, enabling cell to differentiate and adopt the relevant phenotype to reconstruct the ECM. Many parameters are at stake here such as mechanical properties of the scaffold, its structure and alignment as well as chemical cues [102] of the ECM or growth factors. To allow good regeneration, the scaffold should at least be biodegradable or even better, bioresorbable, based on definitions given by Vert [108]. This includes the degradation and the elimination of the polymers and by-products. Time of degradation and the subsequent change in mechanical properties are also crucial for tendon reconstruction but very seldom investigated due to the complexity of such a study. To assess the question, works on small intestinal submucosa (SIS) in canine Achilles tendon repair, carried by Badylak *et al.* (1995) and then Gilbert *et al.* (2007) are valuable. Badylak *et al.* (1995) reported full use of the injured leg by the dogs after 9 to 13 weeks and recovery of injured tendon strength after 12 weeks as assessed by tensile testing [109]. Degradation rate of SIS was investigated by Gilbert *et al.* using carbon 14 measurements, showing 60% loss after 4 weeks and total degradation after 2 months [110]. Thus, we can hypothesize that time span aimed for degradation should be around two months and that, after degradation of the scaffold, recovery and strengthening continue in the month scale.

Last but not least is the processability of the material. This factor is key to meet the specifications listed above and remains one of the main challenges for biomaterials, as the processing should not degrade their properties.

In this review, we will focus on the use of collagen I for tendon reconstruction as it is the main component of tendons and because it is biocompatible and degraded by enzyme with low inflammatory response [111][112]. It also promotes cell adhesion [113][114] and differentiation [102], and is one of the most favorable biomaterials for cell survival compared to cheap and easy to use alginate for instance, or to chitosan [115]. It is thus a promising material that fuels much innovative research [116].

## 5. COLLAGEN BASED TECHNIQUES

As collagen I fulfills the requirements for regenerative scaffolds listed in Table 2, the main challenge is to process it to obtain significant mechanical properties and structure for tendon repair. This section will describe collagen-based techniques, which favor the

production of anisotropic complex architectures, with a particular emphasis on collagen alignment at multiple length scales to better mimic tendon structure. Although promising results for tendon repair have been obtained with isotropic materials such as collagen sponges [117], the focus will be laid on the development of scaffolds which 3D organization aims at providing mechanical support and cell guidance for regeneration.

### 5.1. Extrusion

The extrusion process consists in forcing a material or a solution to pass through a die, giving its shape to the resulting extrudate. This method provides bars, tubes or fibers with a high production rate (few cm/s [118]). Collagen solution extrusion was first developed in the 1960s to create new biosutures for ophthalmic surgeries [119]. Since then, the potential of this molecule was highlighted leading to its widespread use in tissue engineering. Collagen solution extrusion is quite easy to implement requiring only a syringe, a needle or tubing, a collecting bath and a device to control the flow rate (syringe pump) (Fig. 5A). Basically, to produce collagen threads the syringe is filled with a collagen solution and mounted in the syringe pump. Then, the solution will pass through the tubing and will go out in a bath containing the fibrillogenesis buffer. Finally, some other post-production processes can be performed [118], [120]-[126]. Self-assembly of collagen may occur either with a pH transition when using an acidic collagen solution and a basic to neutral pH buffer, or through a temperature jump for cooled neutral collagen solution. In most studies, the collagen is extracted from rat tail (RTT) or bovine Achilles tendons (BAT). Zeugolis *et al.* (2008) compared the properties of threads made of RTT or BAT acidic collagen [121]. By scanning electron microscopy, they observed no significant differences in the substructure of the collagen fibers in the two kinds of threads. Collagen concentration of solution commonly used ranges from 5 mg/mL to 30 mg/mL in acidic conditions (pH=2) [118], [121], [123], [125]-[127]. For the fibrillogenesis buffer, the same basis is found in different studies: 135 mM of NaCl, 30 mM of TES (tris(hydroxylmethyl)methyl-2-aminoethane sulphonic acid) and 30 mM sodium phosphate dibasic with a resulting pH of 7.5 [122], [123], [125], [126]. Other buffers as phosphate saline buffer (PBS) [124] or polyethylene glycol in sodium phosphate dibasic and monobasic [118] are also used. Characteristic dimensions of collagen threads, obtained by this technique, range from 100 µm to 1 mm for wet diameter and around 20 µm to 100 µm after drying [118], [122], [123], [125]. The length depends on the syringe volume capacity and the inner diameter of the tubing. In general, the mean thread length obtained by this technique ranges from few centimeters to about one meter. Mechanical properties of collagen strands have been measured in a routine manner for dry conditions and, less frequently, in wet conditions. Cornwell *et al.* (2007) measured Young modulus of wet collagen fibers around 4 MPa [127]. Regarding the elastic modulus of native tendon, those fibers are between 2 and 3 orders of magnitude “weaker”. For the ultimate tensile strength (UTS), Cavallaro *et al.* (1994) [118], Cornwell *et al.* (2007) [127] and Dunn *et al.* (1993) [125] obtained results ranging from 1.2 to 10 MPa. Dunn *et al.* (1993) also observed that UTS decreases when the fiber diameter increases. So, in wet conditions, which mimics physiological environment, mechanical properties of collagen fibers are still far from that of native tendon (Table 2). This is why different cross-linking (CXL) agents are commonly used to improve those characteristics. Cross-linking process consists in creating new bonds between collagen molecules with chemical species or external excitation sources. The most commonly used on collagen threads are glutaraldehyde, carbodiimide (EDC), cyanamide, hydroxysucrinimide (NHS) and dehydrothermal treatment (DHT), simple drying process, or ultra violet (UV) exposition.

Kato *et al.* (1989) have shown that they can produce collagen threads cross-linked with glutaraldehyde having an UTS of 50 to 66 MPa in wet conditions. Those results are in the same range than
<figure><img src="image_164.png" /><figcaption>**Fig. (5).** Schematics and resulting scaffolds of extrusion (A) and electrospinning (B) techniques. (A) **Top:** Schematic of the extrusion protocol where acidic collagen solution held in the syringe is forced to pass through the tubing and arrives in the fibrillogenesis buffer which induces the thread formation. **Bottom:** collagen thread extruded from an acidic solution at 30 mg/mL observed under optical microscope (left) and under scanning electron microscope (right) (Authors, non-published work). (B) Schematic of the electrospinning protocol where a high voltage is applied between the nozzle and the collector, forming a jet of the polymer solution that dries into fiber. **Inset:** Resulting fibrous mat from a solution in 1,1,1,3,3,3-hexafluoro-2-propanol (Adapted with permission from Bürck *et al.* (2013) [133]. Copyright 2016 American Chemical Society).</figcaption></figure>

UTS of rat tail tendons (33 to 39 MPa). The UTS is divided by a factor 2 for severe dehydration and for EDC treatment. As for Young modulus, they reach 400-500 MPa with glutaraldehyde CXL process. Zeugolis *et al.* (2008) evaluated mechanical properties of collagen threads hung and dried at room temperature [121]. Young modulus of those strands ranged from 1 to 3 GPa and UTS from 0.2 to 0.3 GPa. These results have similar values as native tendon (Table 2). However, we have to keep in mind that in this case, tensile testing is performed in dry conditions, which are not close to the physiological ones. As for Kato *et al.* (1989), they lose on average 3 orders of magnitude in wet conditions. Finally, to have as many bonds as possible, some combinations of CXL processes can be used (dehydrothermal with cyanamide [125] etc).

The alignment of collagen fibrils inside the collagen threads is also a parameter to take into account. The extrusion technique has the main advantage to induce collagen fibrils alignment in the longitudinal axis, thanks to the high constraints applied when collagen solution is forced to pass through the narrow die ([118], [124]). This phenomenon can be further increased by stretching the threads [122].

Lai *et al.* (2011) observed that, *in vitro*, human fibroblasts cultured during 1 or 2 days have an elongated shape following the longitudinal axis of the collagen fibrils [124]. This suggests that there is some contact guidance of the cells on the collagen threads structure. Cornwell *et al.* (2007) highlighted the problem of CXL post-processes on collagen threads. Indeed, they observed that fibroblast migration rate decreased by using CXL agents [127]. A compromise between the improvement of mechanical properties with CXL techniques and the upgrading of cell proliferation must be found. To go further, Dunn *et al.* (1993) implanted their collagen threads in subcutaneous space of the rabbit during 4 weeks. The threads were surrounded by a fibrous tissue and were colonized by inflammatory cells and fibroblasts [125]. Threads crosslinked with DHT combined with cyanamide were degraded over 4 weeks. Cavallaro *et al.* (1994) also had promising results in a canine ligament model and a rat abdominal repair model that can be transposed for tendon repair [118]. They noticed the neovascularization and the cell infiltration in the collagen threads. In both studies, no rejection of the collagen scaffolds was observed.

## 5.2. Electrospinning

Electrospinning is a widespread technique for fibrous scaffold production. This method, based on a simple and inexpensive set-up, uses an electrical field to cast various materials, such as polymer solutions or melts, into fibers, making it attractive for tissue engineering. Briefly, a high voltage is applied between a grounded target and a needle supplied with the liquid of interest, leading to charge accumulation at the liquid surface [128], [129]. For sufficiently high voltage, the surface tension is overcome, leading to the formation of a liquid jet that forms a fiber which dries and is collected on the target, resulting in a fibrous mat (Figure 5B).

To achieve electrospinning of a polymer solution, many requirements must be met. In addition to the setup parameters such as voltage or solution feeding rate, and the ambient conditions such as temperature and humidity, intrinsic characteristic of the solution are paramount to electrospin it. Ideal solutions must have low surface tension and high conductivity [130] and must be viscous enough to form a stable jet that does not break before drying [131] but not too much to ensure sufficient flowability [132] and dry fast before reaching the collector [133]. Usual solvents such as acetic acid lack the viscoelastic properties needed to form a stable jet and do not evaporate fast enough before reaching the collector [133]. Collagen electrospinning was achieved using fluoro-alcohols such as 1, 1, 1, 3, 3, 3-hexafluoro-2-propanol [134]-[136] and 2, 2, 2-trifluoroethanol [137]. However, such a process was proven to denature collagen [133], [138]. Indeed, resulting collagen is soluble in water, does not exhibit characteristic D-banding under transmission electron microscopy and has lower denaturation temperature. Further proofs were given thanks to circular dichroism, electrophoresis and infrared spectroscopy. Attempts succeeded in electrospinning collagen in benign solvent such as Phosphate Buffer Saline and ethanol mix [130] or acidic solution with poly(ethylene oxide) [139], [140]. Bürck *et al.* (2013) demonstrated that collagen fiber electrospun from this solvent did not preserve much better the native structure of collagen and that part of the denatured collagen was intrinsically due to the electrospinning process and not to the solvent [133].

Recently, Elamparithi *et al.* (2015) reported successful electrospinning of collagen in 97: 3 acetic acid: DMSO mix with fibers exhibiting the characteristic D-banding of native collagen [141].
<table><caption>Table 2. Requirements for an ideal scaffold for tendon repair with description and quantitative criteria when assessable.</caption><thead><tr><th></th><th>Requirements</th><th>Descriptions</th><th>Criteria</th><th>Refs.</th></tr></thead><tbody><tr><td rowspan="10">Tendon mimetism</td><td rowspan="3">Size / Shape</td><td rowspan="3">Scaffold size and shape depend on the site of the tendon to be reconstructed</td><td>Rotator cuff tendon</td><td>3 x 21 x 17 mm</td><td rowspan="3">[Verdiyeva 2015]</td></tr><tr><td>Patellar tendon</td><td>5 x 33 x 59 mm</td></tr><tr><td>Achilles tendon</td><td>5 x 14 x 120 mm</td></tr><tr><td rowspan="3">Mechanical Strength</td><td rowspan="3">Can bear usual mechanical sollicitation throughout the healing process and promote cell differentiation</td><td>Young Modulus</td><td>0.5 - 1.9 GPa</td><td rowspan="3">[Ladd 2011, Jung 2009]</td></tr><tr><td>UTS</td><td>50 - 120 MPa</td></tr><tr><td>Strain at Break</td><td>5 - 20 %</td></tr><tr><td>Alignment</td><td>Alignment in the direction of the load to promote cell alignment and differentiation</td><td>Size threshold</td><td>100 nm</td><td>[Van Delft 2008]</td></tr><tr><td rowspan="4">Temporary biocompatible scaffold</td><td>Biotolerable</td><td>Absence of immune rejection from the scaffold and its degradation products in vivo (and non toxic)</td><td colspan="2">Non cytotoxic, genotoxic, carcinogenic ...</td></tr><tr><td>Bioactive</td><td>The scaffold promotes cell adhesion and survival</td><td colspan="2">Must be porous to allow exchange with medium<br>Good colonization and viability of the cells on the scaffold</td></tr><tr><td>Regenerative</td><td>The scaffold fosters the formation of a new tendon by inducing cell differentiation</td><td colspan="2">No observable difference between the new and old tendons</td></tr><tr><td>Biodegradable /resorbable</td><td>Degradation of the scaffold to leave room to the new tendon/and elimination of degradation products</td><td>Time range :</td><td>2 to 3 months</td><td>[Gilbert 2007, Bady-lak 1995]</td></tr><tr><td>Process</td><td>Tunability</td><td>The material can be shaped in a controled way to suit different location and achieve hierarchical structure</td><td colspan="2">Control of the structure from molecular to macroscopic scale, and tunable size and shape of the scaffold</td></tr></tbody></table>

However, no further investigation on native collagen content was carried out. The cross-linking of the resulting fibrous mat after electrospinning with 1-ethyl-3(3-dimethylaminopropyl) carbodiimide hydrochloride does not allow for water solubility test and give surprisingly low mechanical properties with modulus and tensile strength at maximum load of a few kPa. This may be suitable for non-load bearing tissue reconstruction such as cardiac tissue, aimed by the authors, but not for tendon reconstruction.

Thus, electrospinning of collagen for tendon reconstruction seems only relevant when collagen is co-electrospun with biodegradable synthetic polymers such as poly(l-lactide-co-ε-caprolactone) [142] or poly(L-lactic acid) [100], as it improves biocompatibility of synthetic polymers. For instance, an interesting work was published by Ladd et al. (2011) who electrospan two different polymers with collagen to reproduce the musculo-tendinous junction: the poly(L-lactic acid), being stiffer, for the tendon part and poly(ε-caprolactone) for the muscle part [100]. Whereas the synthetic polymers provide mechanical strength, the collagen promotes good cell viability close to 100% after 7 days of both C2C12 myoblasts and NIH3T3 fibroblasts seeded on each part.

### 5.3. Freeze-casting

Freeze-casting, also called ice-templating, is a common technique to produce highly porous scaffolds from solutions or suspensions in solvents such as water or camphene. Considering a water based polymer solution, by simply freezing it, phase segregation occurs between ice crystals and a phase concentrated in polymer (Fig. 6A & B). After freezing, the sample is freeze-dried under vacuum, sublimating ice crystals (Fig. 6B) and results in a dried polymer sponge (Fig. 6C). Depending on the protocol used and the parameters chosen, such as cooling rate, final temperature or size and shape of the vessel, it is possible to favor different crystalline structures and grow ice crystals of different sizes, thus tuning size and shape of the pores [143].

Enabling production of highly porous and anisotropic scaffolds, ice templating of collagen seems promising to promote cell proliferation, alignment and differentiation required for tendon reconstruction. The pores usually obtained in freeze-cast collagen sponges, ranging from 50 to 300 µm, are optimal for cell culture. Switching from small to large pores can be easily done, by decreasing final temperature or cooling rate, or by crystal annealing [144].
<figure><img src="image_165.png" /><figcaption>**Fig. (6).** Freeze-casting setup schematics (A) with the paths followed on the water phase diagram (B) and a resulting collagen sponge observed under SEM (C) and a pre-fibrillated collagen scaffold after freeze-casting (D). (A) Freeze-casting schematic where a collagen solution is directionally frozen on a metallic cooled plate, which other side is dipped in liquid nitrogen and which temperature is controlled thanks to a thermocouple and a heating coil; causing segregation between ice crystals and a concentrated phase in collagen. (B) Water phase diagram schematic where the path followed during freeze-casting (1) and then freeze-drying (2) is drawn. (C) An isotropic collagen sponge observed under SEM (Adapted from Davidenko et al. 2015 [150], Creative Common License). (D) SEM picture of Lowe et al.'s longitudinal scaffold, pre-fibrillated before freeze-casting in a high aspect ratio vessel (Adapted with permission from Lowe et al. 2016 [151]. Copyright 2016 American Chemical Society)</figcaption></figure>

Even though the principle of freeze-drying seems simple, the physics behind is not straightforward. Pawelec and co-workers have realized interesting observations on collagen solution freeze-casting and the dependence of pore size [145] and anisotropy [146] upon parameters. They also investigated the effect of the addition of NaCl and sucrose [147], and of heat transfer between the mold and the heat sink [148] on collagen freeze-casting. They took their work one step further by culturing ovine primary patellar tendon cells on freeze-casted collagen uniaxial and anisotropic sponges in a fibrin gel [149], exhibiting higher cell proliferation, metabolism and matrix production thanks to fibrin gels and a cell phenotype control by the collagen scaffold structure.

Whereas Pawelec and coworkers mainly focused on the freeze-
casting of the collagen itself, Caliari and coworkers rather investi-
gated cell culture on freeze-casted scaffolds for tendon reconstruc-
tion. Using collagen mixed with glycosaminoglycan (10: 1) (Coll-
GAG), Caliari and Harley demonstrated that pores above 150 µm
were more favorable for cell proliferation and metabolism [152]
while a mix of Insulin Growth Factor 1 (IGF-1) with Growth/ Dif-
ferentiation Factor 5 (GDF-5) was more effective to promote cell
proliferation, collagen production and tendon associated structural
protein gene expression [153]. Given the relatively low mechanical
properties of collagen sponges (modulus around 800 kPa in dry
state), Caliari et al. (2011) made an attempt to compensate this by
wrapping the freeze-casted scaffold in a collagen air dried mem-
brane crosslinked with 1-ethyl-3-[3-dimethylaminopropyl] carbodi-
imide hydrochloride and N-hydroxysulfosuccinimide, resulting in a

core shell scaffold with a 8 MPa modulus, UTS between 0.3 and 0.9
MPa and strain at break from 10 to 25 % [154]. Ice templating be-
ing also promising for bone reconstruction research, they also pro-
duced a bi-component material Coll-GAG/ Calcium phosphate
mineralized Coll-GAG (CaP Coll-GAG) scaffold to mimic bone-
tendon junction, together with specific growth factors on each side
to study the impact on mesenchymal stem cells (MSCs) [155]. Each
side exhibited up-regulation of pro-osteogenic and pro-tenogenic
genes in the CaP Coll-GAG and Coll GAG parts respectively.

In most articles, two points are not tackled. The first and more
important one is the self-assembly of collagen, which could im-
prove mechanical properties, cell adhesion and differentiation
[113], [114]. The second one is the shape of the scaffolds that dif-
fers from tendons. Since the geometry of the vessel and its filling
height highly impact the scaffold structure, any modification of the
former will impact the latter. This increases the difficulty to mimic
living tissues with this method. However, Lowe et al. addressed
those two problems [151]. By ice-templating self-assembled colla-
gen, they produced high aspect ratio and highly aligned porous
collagen scaffolds (Figure 6D) that could be suitable for Achilles
tendon for instance. Uniaxial micro features at the surface enabled
rat dermal fibroblasts and chick dorsal root ganglia explants align-
ment. Precise pore analysis was not reported, neither cell culture
inside the scaffold. It should be noted that the scaffolds were ob-
tained without any cross-linkers, and that even though the mechani-
cal properties of the scaffold are low (Young modulus ~600 kPa,
UTS 150 kPa, strain at break 110%), they are in the same range as
those obtained with cross-linked but not fibrillated scaffolds [154].
The addition of carbodiimide cross-linker could further stiffen the
material.

On the way to improve mechanical strength and modulus of scaffolds, an innovative design was developed by Mozdzen et al. [156]. They freeze-casted a collagen-GAG solution into a 3D printed acrylonitrile butadiene styrene (ABS) array. Even though ABS may not be the most suitable polymer for tendon reconstruction because of its lack of biocompatibility [157], the composite design of this scaffold is of interest. It combines the mechanical strength of ABS with the high biocompatibility of collagen sponges. Comparing sinusoidal ABS fiber arrays of different amplitude to straight ABS fiber arrays, they were able to tune elastic properties of the scaffold in a crimp-like fashion. This gives a glimpse of the promises held by 3D printing for tissue reconstruction.

5.4. 3D Printing

3D printing, or additive manufacturing, has breached in most
good manufacturing sectors in the last few years and tissue recon-
struction is no exception. Based on the knowledge acquired on scaf-
fold manufacturing and facing new intrinsic problems, it gives high
hopes for its capacity to design complex macroscopic shapes of
scaffolds as seen in the previous paragraph, as well as enable pre-
cise cell deposition. It consists in a layer-by-layer computer aided
deposition that enables the construction of 3D structure designed on
the computer.

This technique is currently used in research on skin, cartilage,
bone, aortic valve [158]. Different methods of deposition have been
developed such as inkjet, extrusion (Figure 7A left) or laser-assisted
bioprinting [158]-[160], as well as different hydrogel based inks,
called bio-inks [115]. The use of collagen as an ink for 3D printing
raises an obvious interest and is investigated especially for bone
[161]-[163] and cartilaginous tissue reconstruction [164], [165].
However, current research face difficulties to obtain self-standing
3D printed structures. Thus, collagen is often used to enhance the
bioactivity of widespread bioinks such as alginate [166], [167] or
agarose [162] that are cheap and easy to use but show lower bioac-
tivity compared to main components of the ECM. It is also used
with calcium phosphate [161] for bone reconstruction or mixed
with fibrinogen [168] that is easy to crosslink.
<figure><img src="image_166.png" /><figcaption>**Fig. (7).** **Schematics and resulting scaffolds of extrusion 3D printing and electrochemical techniques** (B). (A) **Left:** schematic of 3D printing setup where a dispensing unit is monitored in the three directions of space by a computer to deposit bio-ink layer by layer. **Top right:** 3D printed scaffold to reproduce myo-tendinous junction, accurately seeded with DiO-labeled C2C12 myoblasts (green - top) and DiI-labeled NIH/3T3 fibroblasts (red - bottom) after 1 day in culture. **Bottom right:** Same scaffold after 7 days in culture exhibiting the interface region in yellow. (B) **Top:** schematic of the electrochemical technique where a voltage is applied to a collagen solution between the anode and the cathode, creating a pH gradient and inducing collagen migration to the isoelectric point (pI) region and alignment of the molecules. **Bottom left:** picture of a thread of electrochemically aligned collagen (ELAC) between the two electrodes observed with polarized microscopy. **Bottom right:** picture of a scaffold woven from triple ELAC threads twisted together. (A, top and bottom right) adapted with permission from Merceron *et al.* (2015) [176]. (B, bottom) adapted with permission from Younesi *et al.* (2015) [179].</figcaption></figure>

Other techniques to print collagen self-standing structures are the deposition of collagen on a cold plate to freeze the solution when deposed [169], [170], or the use of sacrificial negative molds [171], [172]. An original process that is worth mentioning is the printing of a core-sheath collagen-alginate structure [173].

Here again, collagen is sometimes used without inducing fibril-
logenesis. Besides, the protocols used can be too harsh to incorpo-
rate cells in the bio-ink, preventing the use of additive manufactur-
ing to deposit cells in an accurate and homogeneous way. A few
articles deal with pure collagen self-assembled during or after print-
ing [164], [174]. However, for collagen [174] as well as for fibrino-
gen [168], this leads to tedious protocols where the scaffold is incu-
bated, after each layer printing, either at 37°C or with thrombin
respectively.

One currently adopted strategy is the printing side-by-side of an
artificial biodegradable polymers and a bio-ink with incorporated
cells. The artificial polymer confers mechanical integrity to the
scaffold whereas the bio-ink allows exact cell deposition with suit-
able bioactivity [168], [175]. To our knowledge, despite the control
on porosity, macroscopic structure and cell incorporation that this
technique offers, no article about 3D printing of collagen for tendon
reconstruction has been published yet. This can be explained by the
difficulty of handling collagen and the poor mechanical properties
obtained with collagen in other domains such as extrusion. As col-
lagen is often used with other bio-inks for its bioactivity, some
chose gelatin instead for its lower price. For instance, Merceron *et
al.* (2005) published the co-printing of two synthetic polymers
polyurethane (PU) and poly(ε-caprolactone) (PCL) together with a
bio-ink based on hyaluronic acid, fibrinogen and gelatin, to repro-
duce the musculo-tendinous junction [176]. The bio-ink contained
C2C12 myoblasts on the PU side while the bio-ink on the stiffer
PCL side contained NIH/3T3 (Figure 7A top right). The construct
exhibited good cell viability (Figure 7A bottom right), high cell
alignment and characteristic musculo-tendinous gene upregulation.
They highlighted the ability offered by 3D printing to deposit dif-
ferent cell types in precise locations, paving the way to complex
multicellular scaffolds for regeneration.

**5.5. Electrochemical Techniques**

Applying an electric field to a collagen I monomer solution
enables to control the spatial aggregation of molecules, resulting in
collagen membranes [177] or bundles measuring 50 to 400 µm in
diameter and several centimeters in length [178]. Electrochemical
reactions primarily due to the electrolysis of water create a pH gra-
dient. In the pH region corresponding to the isoelectric point, the
collagen becomes less sensitive to the electric field and stops mi-
grating due to global neutral charge (Figure 7B top and bottom left,
[179]). In these biomimetic constructs small fibers are highly ori-
ented along the bundle axis, increasing the mechanical properties as
compared to randomly oriented controls, and inducing an alignment
of tendon derived fibroblasts that migrate into braided bundles
[178]. Isoelectric focusing technology was further adapted to en-
hance directional neuronal growth [180]. This study also unraveled
the underlying hierarchical structure, with nanofibrils aggregating
into ~500 nm diameter D-banded fibrils that align to form larger
entities. The influence of phosphate buffer and incubation time
on strength and D-banding pattern of the electrochemically aligned
collagen (ELAC) threads was also studied and advocated PBS 1X
during 12h as the optimal fibrillation buffer [181]. After 96h incu-
bation, Young modulus could attain 6 MPa in wet conditions. The
ability to induce tenogenic differentiation of human MSCs was
evidenced on the threads with the increase of tendon specific mark-
ers such as scleraxis and tenomodulin [182]. An in vivo study was
led to assess the response of rabbit patellar tendon (PT) to braided
ELAC threads [183]. Four months post-implantation, cross-
sectional area, mechanical properties and tendon-fascicles content
of the PTs treated with ELAC were optimized as compared to con-
trols, but these differences were not evident anymore after 8
months. A low-grade granulomatous inflammation was observed at
4 months but it spontaneously diminished between 4 and 8 months.
Different crosslinking strategies were investigated to improve the
mechanical properties of ELAC threads, and the increase of genipin
concentration in conjunction with highly concentrated ethanol was
advocated [184]. Further processing was made with the fabrication
of collagen textiles (Fig. 7B bottom right) by weaving ELAC triple
threads after twisting continuous ELAC threads obtained with a
kinematically rotating linear electrode pair [185]. MSCs seeded in the porous scaffolds underwent tenogenic differentiation, and the mechanical properties reproduced some aspects of the functional mechanics of native tendon, in particular comparable load-displacement curves in the linear regions. Computer aided design that helped produce patterned electrodes enabled to obtain highly porous collagen scaffolds with controlled geometries that can be further combined to form layered structures [186]. The elastic modulus of these different types of constructs ranged from 5-40 MPa.

## 5.6. Cell-mediated Collagen Gel Compaction

Cells embedded in loose collagen hydrogels have the ability to compact them into denser tridimensional constructs. This cell-mediated collagen gel contraction was first reported in 1979 by Bell and coworkers [187]. Among the first investigators studying this phenomenon, Stopak and Harris showed that it stems from cell contractility [188]. They also embedded fibroblastic cell explants in collagen gels and observed both fiber alignment and cell orientation along the axis between the explants. They carried out other experiments in which fibroblasts, initially dispersed within a gel, could contract the matrix between fixed posts (Fig. 8A). This resulted in complex patterns of crisscrossing strands. In order to explain the reorientation of the different components of cellularized gels Barocas and Tranquillo proposed in 1997 a mathematical model in which fibril alignment results from traction forces exerted by the cells under anisotropic conditions, such as a mechanical constraint or inhomogeneous cell distribution. According to this model, cells then align along the fibrils by contact guidance [189]. Huang et al. (1993) showed an increase in stiffness over time of fibroblast-populated collagen constructs between fixed posts, and demonstrated that lysyl-oxidase crosslinking activity was necessary for this mechanical enhancement [190]. Autologous bone derived MSCs were further used for Achilles tendon repair in rabbits [191]. In these experiments Young et al. (1998) used a cell-gel composite contracted onto a pre-tensioned suture (Fig. 8B). They showed that the mechanical properties were greatly enhanced 12 weeks after implantation in 1-cm-long gastrocnemius tendon gaps compared to controls implanted with suture material only. MSC-collagen composites, implanted into central patellar tendon defects of rabbits, showed a significant improvement of the biomechanical properties as compared to natural repairs, with cells remaining within the repair site [192], [193]. No additional benefit of increasing cell density, from 1 to 8 million per ml, was observed. Formation of ectopic bone in 28% of the composite grafted repairs was noticed. Lowering the cell to collagen ratio in a different system, where the MSCs were placed into silicon dishes between fixed posts 2 weeks prior to implantation (Fig. 8A), showed no significant difference in material and histological properties between implants with 40, 000 cells/mg collagen or 80, 000 cells/mg collagen twelve weeks after surgery [194]. There was also no significant difference in mechanical properties between these implants and acellular ones, with an average modulus reaching 20% of normal patellar tendons.

Scaffold-free constructs were also obtained by self-assembly of collagen and non-collagenous molecules produced by rat Achilles tendon cells grown to confluence between two pinned sutures, serving as anchor points, for 2 weeks [195] (Fig. 8C). The obtained cylinders presented a tangent modulus of 17 MPa, as compared to 27 MPa for embryonic chicken extensor tendons. Tendon-skeletal muscle constructs were also engineered in vitro in scaffold-free environment, with the reconstitution of myo-tendinous junctions either by co-culturing fibroblasts and myoblasts between fixed posts [196] or by pinning tendon constructs on muscle cell monolayer [197]. Fibrin has also been used as a material for tendon engineered constructs. Kapacee et al. showed that in fixed length fibrin gels made with tendon embryonic cells, fibrin was replaced by

<figure><img src="image_167.png" /><figcaption>**Fig. (8).** Schemes of various types of tendon constructs based on cell-mediated gel compaction. On these top-views cell-gel mixtures are depicted pink, and the contraction process is represented over time, from top to bottom. (A): Principle of collagen matrix contraction between fixed posts, depicted white, as introduced in [187] and further developed in [188], where cells and collagen gels are first pipetted into wells. The cells contract the gels around two posts located at the base of each well. (B): Collagen gel-cell composite contracted onto a pre-tensioned suture, as introduced in [191]. Each construct was created in a glass trough, which contained a tensioned suture (horizontal arrows). (C): Tendon construct formation between pinned sutures, in the presence of cells only [195], fibrin-cell mixture [198], or collagen-cell mixture [201].</figcaption></figure>

aligned collagen fibrils within 6 days [198]. Similar tendon constructs were also used to investigate the role of transcription factor EGR1 towards tendon differentiation by means of forced Egr1 expression programmed MSCs in a fibrin gel [199]. Neal et al. (2014) developed a method using a PDMS mold and a second sacrificial mold, made of gelatin, and placed inside, to produce thin fascicle-like constructs, ~100 µm in diameter, of fibrin hydrogel with cells embedded in it [200]. This strategy enabled to combine molding and cell-mediated gel compaction of the freely suspended hydrogel anchored to the PDMS walls. C2C12 mouse myoblasts alignment and maturation were promoted. According to preliminary work, the method could also be extended to 3T3 fibroblasts and horse tenocytes. More recent studies, led by two different groups, showed that fibrin gels enhance the properties of tendon-engineered constructs over collagen-based materials. Yeung et al. (2015) led a comprehensive study of chick tendon fibroblast transcriptomes inside gels between pinned sutures and showed that fibrin-based constructs recapitulated more closely gene expression that occurs during tendon development as compared with collagen-based ones [201]. Cell-synthesized collagen matrix also exhibited better fibrils alignment in the fibrin-based gels. Breidenbach et al. (2014) showed that for approximately 14 days in cell-gel mixtures between posts embyro tendon/ligament progenitor cells exhibited enhanced tenogenic expression, collagen alignment and linear modulus with fibrin [202]. Still, a reduction in mechanical, biological and mechanical properties was observed in fibrin constructs submitted to cyclic stretching (2.4% strain, 1Hz, 3000 cycles/day) whereas this was beneficial to collagen constructs. The main conclusion is that the various parameters of mechanical stimulation must be fine-tuned for each type of construct.

## 5.7. Other Promising Methods

In this section we will describe some other methods, which have been applied to the generation of a high anisotropy inside collagen solutions. They often rely on the combination of multiple physical phenomena, and adapted for a better processability they may be promising candidates to generate biomaterials for tendon repair.

### Magnetic Alignment

Torbet and Ronzière were the first to propose the magnetic alignment of collagen during self-assembly in solutions at low concentration [203]. They used magnetic field strengths between 1.9 and 5.6 T. Aligned fibrils being in the same plane as the field, they
inferred the additional ordering effects of surface interactions. Guido and Tranquillo further showed that fibril orientation could be controlled by independently varying the magnetic field strength or temperature during fibrillogenesis, pointing out the possible orienting effect of the magnetic field acting over an extended fibrillogenesis period due to a lower temperature [204]. When human foreskin fibroblasts are cultured in the gel, their orientation correlates with that of the fibrils. Chen *et al.* (2011) showed that, in tilapia fish-scale collagen solutions, fibril alignment can also be induced after fibrillogenesis with magnetic fields of 6 or 12 T [205]. The rheological properties obtained were similar in both cases, with either non-prefibrillated or fibrillated and crosslinked collagen prior to magnetic treatment, with higher values than those of un-oriented gels. Those properties were dependent on strength and application time of the magnetic treatment. In both cases, collagen fibrils maintained a typical D-periodic structure of 60-70 nm. Guo and Kaufman (2007) showed that it was also possible to obtain fibril alignment with a small magnet by combining flow and magnetic field effect [206]. Surface modified paramagnetic iron oxide beads placed inside collagen solutions induce collagen alignment within areas of hundreds of square microns, the domains being larger when timescales of gelation and bead motion to the magnet poles are similar. Multiple layers of collagen are obtained. C6 glioma cells embedded in the matrix were able to remodel it by bundling the fibers.

### Microfluidic alignment

Lee *et al.* (2006) proposed the microfluidic alignment of collagen fibrils in dilute solutions under flow, and the orientation obtained appeared to be particularly efficient within PDMS microchannels 10 to 100 µm wide [207]. They inferred that the rapid attachment of fiber precursors along the channel walls and the subsequent influence on new fiber growth were involved, but mentioned that no firm conclusion could be drawn from their experiments to provide an explanation. Aortic bovine endothelial cells grown on the UV-crosslinked collagen matrix aligned along the direction of the fibers. Saeidi *et al.* (2009) studied the formation of fibrils on glass surface under the influence of flow between two plates [208]. The height of the microchamber could be changed which enabled to vary the shear rate at the walls. In this context self-assembly process is influenced by shear rate, as well as self-assembly energetics and surface energy. Moderately aligned collagen fibrils were obtained. They result from the aggregation of 3-5 nm microfibrils and present no D-banding, possibly due to the distortion of the fibrillar structure by the surface energy of the glass. The best alignment was observed at shear rates between 20 and 80 s⁻¹, and above these values the fibrils would often turn down-stream due to instabilities. Surface bound aligned collagen I matrices were produced on copolymer-coated or glass substrates thanks to a microfluidic system [209]. Collagen deposited under flow was either pre-fibrillated or not. The morphology of the matrices obtained could be controlled. Fibril coverage was directly dependent on collagen concentration, with extended matrices at 0.8 mg/ml as compared to 0.2 and 0.4 mg/ml. The flow rate also had a direct impact on the alignment of pre-fibrillated collagen, with highest alignment degree at 11 µl/min as compared to 4 and 0.45 µl/min. A slight decrease of alignment was seen on more hydrophilic surfaces. Pre-conditioning of non-fibrillated collagen (0.8 mg/ml) in heated tubing before entering the channel had a direct impact on the fibers obtained, which are longer and more individual after 30 minutes as compared to 5 and 10 minutes.

Recently Haynl *et al.* (2016) used a microfluidic chip connected to three syringes to produce collagen microfibers upon an increase in the pH and the presence of PEG [210]. The diameters of the fibers can be varied depending on the extrusion buffer and the flow rates. With a collagen flow rate of 50 µl h⁻¹ and buffer flow rates in between 235 and 550 µl h⁻¹ the fibers could be collected using a rotating spool. The diameters obtained under these conditions are

around 10 µm and after air-drying from 3 to 6 µm. Alignment of the fibrils within the microfibers was evidenced by scanning electron microscopy and polarized FTIR spectroscopy. Young moduli obtained were 4138 +/-512 MPa after drying. Neuronal cells cultured on the microfibers aligned along the longitudinal axes.

### Rotary shearing

Yunoki *et al.* (2015) produced thick collagen gel bundles with aligned fibrils by means of rotary shearing to a dense collagen solution (1.8%), with gelation triggered upon rotation [211]. Well-aligned fibrils, as evidenced by scanning electron microscopy and birefringence measurements, were observed in the marginal regions of 1-3 mm thick disc-shaped gels. The optimal shear-rates for fibrils alignment were between 2-5 s⁻¹. Rectangles and dumb-bell-shaped specimens cut from the discs (7\*12mm and 4\*16mm respectively) in the direction of the rotation had Young modulus of 180 kPa after crosslinking with genipin, as compared to 30 kPa for the rectangles perpendicular to the rotation. After EDC/NHS crosslinking the values reached were 280 and 130 kPa, respectively. 3T3 cultured on aligned matrices were spindle-shaped in the fibril alignment direction.

### Imprinting

An innovative technique was invented by Tong *et al.* to study the role played by the biophysical and biochemical environment, as well as mechanical properties, on MSCs differentiation into tenocytes [102]. They replicated tendon structure by imprinting longitudinal sections of bovine Achilles tendons with PDMS, making a negative then a positive replica reaching sub-micron precision (Fig. 9).

<figure><img src="image_168.png" /><figcaption>**Fig. (9).** Schematics of the imprinting technique. A PDMS negative replica of a longitudinal tendon cryo-section is made to produce a subsequent PDMS positive replica.</figcaption></figure>

By changing monomer concentration, they were able to tune the elastic modulus of the imprint from 0.35 to 3.5MPa. They demonstrated that surface topography, collagen coating, and stiffness were paramount to promote MSCs differentiation. In addition to such valuable information, this technique could be used to mold scaffold for tendon reconstruction, resulting in optimal surface topography.

### Fiber drawing

Another interesting, yet simple technique, has been reported by Paten *et al.* (2016) [212]. They were able to draw centimeter-long collagen fibers with a diameter around 50 µm, just by poking a collagen solution drop left a few minutes under dry nitrogen atmosphere with a glass micro-needle (Fig. 10). The flow-induced crystallized fibers presented outstanding aligned structure on the outer
shell. Fibers were stable when kept under strain or in polyethylene glycol solution. This could help understanding tendon morphogenesis and, above all, by weaving such fibers, it could provide a promising and simple scaffold production, with inherent high porosity and alignment to promote cell proliferation and differentiation.

<figure><img src="image_169.png" /><figcaption>**Fig. (10).** Schematics of the fiber drawing technique. A glass micro-needle is used to poke the surface of a collagen solution left 150s to concentrate under nitrogen atmosphere, and draw a fiber.</figcaption></figure>

### Plastic compression and gel aspiration-ejection

Plastic compression is a cell-free, controlled process first introduced by Brown *et al.* [213], to produce dense, cellular, mechanically strong native collagen structures. This technique relies on external mechanical loading and capillary fluid flow to expel interstitial liquid. The cell viability is high during and after compression of cellular collagen gels, even surpassing highly hydrated collagen gel counterparts, with an increase in proliferation over time [214]. Plastic-compressed collagen gels seeded or not with rabbit flexor tendon cells, 20-40 µm thick, were rolled in order to produce spiral rolls 22 mm in length x 2.25 mm in diameter [215]. These rolls were placed in a test site across intercostal spaces in rabbits to provide tensile cyclic loading *in vivo* for 5 weeks. Collagen remodeling and tensile mechanical strength were enhanced in the cell-seeded gels. Gel aspiration-ejection (GAE) technique was further developed to produce anisotropic, cell-seeded, injectable collagen, without the use of crosslinking [216]. After incubation of low concentrated collagen solutions the resulting gels are aspirated into capillaries by means of a negative pressure applied with a syringe. Fibril alignment was evidenced by scanning electron microscopy and polarized ATR-FTIR spectra. 3T3-seeded collagen gels submitted to GAE showed a rapid orientation of the cells along the direction of the fibrils within one day. Constructs obtained through this process and exposed to the appropriate supplements also accelerated the differentiation of seeded murine MSCs towards osteoblastic and neuronal lineage, as compared to plastic-compressed cell-seeded constructs.

## 6. DETERMINANT IMPACT OF MECHANICAL STIMULATION

Tendon development, homeostasis and healing are influenced by their loading environment [2]. During rehabilitation, a fine balance between complete immobilization and loading is necessary to improve ECM production and tendon repair [217]. Before implantation, engineered constructs can be exposed to static and dynamic loads in order to better mimic the *in vivo* conditions and potentially improve their properties. The results obtained may also give in-

sights into the influence of mechanical stimulation on cell differentiation, activity and on biomechanical enhancement (Table 3).

The influence of uniaxial loads on ECM components was studied using collagen constructs without cells. Static stretching of self-assembled collagen fibers extruded in phosphate buffer, then air dried and rehydrated, induced a modulus increase from 1.82 MPa with 0% stretch to 45 MPa after 50% stretch, measured after air drying [218]. Increased collagen fibril orientation was supposedly involved based on birefringence values. Static stretching of PDMS substrates coated with a multilayer film composed of collagen and alginate enabled to produce surfaces with aligned collagen fibrils, that were further stabilized with genipin crosslinking, with an optimal strain of 100% for alignment [219]. Cyclic loading of strips of collagen gels obtained by plastic compression promoted fibril fusion as evidenced by transmission electron microscopy [220]. Tensile tests showed a modulus increase from 244 kPa after 1 cycle to 535 kPa after 144 cycles, with 20% maximum strain for each cycle. There was also an increase in break strain and a 4.5 fold rise in break stress. Regarding the mechanical stimulation of cells alone, a cyclic axial strain applied on human tendon fibroblasts showed a strong influence on cell proliferation, highlighting the need to fine-tune the stretch [221].

Continuous cyclic strain inhibited proliferation of four types of cells including rabbit flexor tendon derived cells, whereas an intermittent cyclic strain could enhance cell proliferation in adipod-derived stem cells and sheath fibroblasts [222]. Rat bone marrow MSCs submitted to an uniaxial cyclic stretching of 10% at 1Hz for durations ranging from 3 to 36 h showed an increased expression of collagen types I and III and of tenascin-C [223]. The percentage of cells oriented along or close to the stretch axis was decreased, as previously noticed with ligament fibroblasts [103] as well as with epitenon tenocytes [222]. The reorientation of the different components of cellularized gels under mechanical stimulation thus appears complex.

Garvin *et al.* imposed a uniaxial strain on bioartificial tendons made of avian flexor tendon cells in a collagen gel, with an elongation pattern of 1% at 1 Hz for one hour each day [224]. Results obtained in the first week of culture showed an alignment of the cells along the direction of strain and a nearly 3-fold increase in ultimate tensile strength as compared to the non-loaded constructs. Human bone-marrow derived MSCs in a collagen gel were also submitted to a similar stimulation protocol for 30 minutes each day [225]. Expression of scleraxis, the main tendon marker during vertebrate development, was up-regulated in the 3D constructs as compared to 2D culture, but this was maintained over time only in the constructs submitted to cyclic loading. The observed tendinous neo-tissue development could be associated with matrix remodeling activity, in agreement with the noticed modulation of matrix metalloproteinases expression. Another study led by Scott *et al.* (2011) with multipotent mesenchymal cell line C3H10T1/2 in the same 3D model optimized the protocol of stimulation based on the expression of scleraxis and type I collagen [226]. The insertion of a 10 s resting period between each repetition of stretching cycle (10% strain, 0.1 Hz) appeared to be a critical parameter.

Mechanical stimulation of another type of scaffold based on collagen type I sponge, with rabbit bone marrow MSCs applied on it, was also investigated prior and after surgical implantation into patellar tendon central third defects [227], [228]. When the peak strain was 2.4%, applied every 5 minutes for 8 hours each day for two weeks, the stimulated constructs had 4 times the linear modulus and 2.5 times the linear stiffness of their non-stimulated counterparts *in vitro* [228].

This was associated with increased expressions of collagen type I and type III. Repair tissues formed with constructs stimulated 2 weeks with 4% strain prior to implantation showed improved
<table><caption>Table 3. Mechanical stimulation protocols and main results reported for cells alone, collagen constructs, and mixtures of cells and collagen.</caption><thead><tr><th>Scaffold</th><th>Cells</th><th>Stimulation</th><th>Main results</th><th>Refs.</th></tr></thead><tbody><tr><td>Collagen thread</td><td></td><td>Static axial strain for 24h</td><td>Elastic modulus: 45 MPa after 50% stretch / 1.82 for the controls (air-dried).<br>Increased axial fibril orientation (birefringence)</td><td>[Pins 1997]</td></tr><tr><td>Strips of collagen gels obtained by plastic compression</td><td></td><td>Cyclic loading<br>20% strain maintained 5 min for each cycle (20 min total)</td><td>Elastic modulus: 535 kPa after 144 cycles / 244 kPa after 1 cycle.<br>UTS: 4.5-fold increase</td><td>[Cheema 2007]</td></tr><tr><td>PDMS substrate coated with multiple layers of collagen/alginate</td><td></td><td>Static stretching</td><td>Fiber alignment in the stretching direction with optimal strain of 100%.<br>Cell alignment with the fibrils</td><td>[Chaubaroux 2015]</td></tr><tr><td rowspan="2">Rabbit flexor tendon derived cells /Adipoderived stem cells /Bone marrow derived stem cells</td><td></td><td>Continuous cyclic strain 8% - 1 Hz</td><td>Proliferation inhibited</td><td rowspan="2">[Riboh 2008]</td></tr><tr><td></td><td>Intermittent cyclic strain 4% - 0.1 Hz<br>On/off: 1h /2h or 1h/5h</td><td>Enhanced proliferation and collagen expression.<br>Cells perpendicular to the strain axis</td></tr><tr><td>Rat bone marrow MSCs</td><td></td><td>Uniaxial cyclic stretching 10% - 1 Hz<br>Duration: 3 to 36h</td><td>Increased expression of collagens type I and III and tenascin-C.<br>Cell reorientation with less cells along the stretch axis</td><td>[Zhang 2008]</td></tr><tr><td>Collagen gel</td><td>Avian flexor tendon cells</td><td>1% - 1 Hz<br>1h/day</td><td>Alignment of the cells along the direction of strain.<br>UTS: 3-fold increase compared to controls</td><td>[Garvin 2003]</td></tr><tr><td>Collagen gel</td><td>Human bone marrow MSCs</td><td>1% - 1 Hz<br>30 min/day</td><td>Scleraxis up-regulation maintained with cyclic loading<br>Regulation of matrix metalloproteinases expression</td><td>[Kuo 2008]</td></tr><tr><td>Collagen gel</td><td>Mesenchymal cell line C3H10T1/2 (mouse)</td><td>Variable strain -0.1 Hz</td><td>Increased expression of scleraxis and collagen type I with 10% strain.<br>Further increase when a 10s resting period is inserted between each cycle repetition</td><td>[Scott 2011]</td></tr><tr><td>Collagen sponge</td><td>Rabbit bone marrow MSCs</td><td>4% - 1Hz<br>cycle every 5 min<br>8h/day</td><td>2.5-fold rise in linear stiffness in vitro<br>12 weeks after in vivo implantation into patellar tendon central third defects: maximal forces sustained are 70% of normal tendons / 55% for non-stimulated constructs</td><td>[Juncosa-Melvin 2006]</td></tr><tr><td>Collagen sponge</td><td>Rabbit bone marrow MSCs</td><td>2.4% - 1Hz<br>cycle every 5 min - 8h/day</td><td>4-fold rise in linear modulus and 2.5-fold rise in linear stiffness in vitro<br>Increased expressions of collagen type I and type III</td><td>[Juncosa-Melvin 2007]</td></tr><tr><td>Collagen sponge</td><td>Rabbit bone marrow MSCs</td><td>Variable strain - 0.1Hz<br>Variable cycle number and cycle repetition</td><td>Constructs stimulated with 2.4% strain, 3000 cycles/day, and one cycle repetition produced the stiffest constructs, peak strain appearing as the most critical parameter</td><td>[Nirmalandhan 2008]</td></tr><tr><td>Collagen gel combined with collagen sponge</td><td>Rabbit bone marrow MSCs</td><td>2.4% - 1Hz<br>cycle every 5 min - 8h/day</td><td>12 weeks after in vivo implantation into patellar tendon central third defects: enhanced mechanical properties in the repair tissue obtained with stimulated constructs</td><td>[Shearn 2007]</td></tr></tbody></table>
<table><caption>Table 4. Summary of experiments and main results obtained using newly developed collagen-based scaffolds in animal models.</caption><thead><tr><th>Technique</th><th>Scaffold/parameters</th><th>Animal models / Sample size</th><th>Implantation site / Controls</th><th>Main results</th><th>Refs.</th></tr></thead><tbody><tr><td rowspan="3">Extrusion</td><td>Parallel crosslinked collagen fibers (200) in an uncrosslinked collagen matrix<br/>-Fiber diameter: 20 or 50 µm<br/>-Cross-linking: GLUT or DHTC</td><td>Rabbit<br/>n=5 for each group</td><td>Subcutaneous</td><td>After 4 weeks:<ul><li>DHTC-crosslinked fibers mostly degraded and replaced by fibrous tissue</li><li>GLUT-crosslinked fibers mostly intact, surrounded by inflammatory cells</li><li>No influence of initial fiber diameter</li></ul></td><td>[Dunn 1993]</td></tr><tr><td>Knitted collagen fabrics<br/>-Dry diameter: 25 µm<br/>-Cross-linking: EDC</td><td>Dog</td><td>Anterior Cruciate Ligament</td><td>After 12 weeks:<ul><li>Applied force by the operated limb at 70% of control</li><li>Threads remain intact with disorganized cellular ingrowth</li><li>Neovascularization, normal size collagen fibrils in the neoligament</li></ul></td><td>[Cavallaro 1994]</td></tr><tr><td></td><td>Rat<br/>n=5</td><td>Abdominal wall defect</td><td>After 3 weeks:<ul><li>Neovascularization and fibroblastic cellular infiltrate</li></ul>After 12 weeks:<ul><li>No herniation</li></ul></td><td></td></tr><tr><td>Electrochemical techniques</td><td>Braided ELAC threads (9)<br/>-Threads diameter: 200-300 µm thick, 400-500 µm wide<br/>-Cross-linking: Genipin</td><td>Rabbit<br/>Sacrificed at 4 and 8 months (n=3 each)</td><td>Patellar tendon (incised along the length)<br/>Controls: incised contralateral knees</td><td>After 4 months:<ul><li>Low degradation of the ELAC scaffold</li><li>Cross-sectional area, mechanical properties and tendon-fascicles content of the PTs treated with ELAC optimized / controls</li><li>Low-grade granulomatous inflammation</li></ul>After 8 months:<ul><li>Differences / controls less evident</li><li>Diminished inflammation</li></ul></td><td>[Kishore 2012]</td></tr><tr><td>Cell-mediated gel compaction</td><td>Bone derived MSCs-collagen gel composite - Contraction: pre-tensioned suture</td><td>Rabbit<br/>Sacrificed at 4, 8, or 12 weeks (n=16 each)<br/>5 animals used for establishing normal biomechanical properties</td><td>Achilles tendon (1 cm long gap defect)<br/>Controls: contralateral tendons implanted with suture material only</td><td>After 4 weeks:<ul><li>Stiffness is 1/2 and maximum force 2/3 of normal tendons, 2X / controls</li></ul>After 12 weeks:<ul><li>Modulus 1/2 of normal tendons, 2X / controls</li><li>Histology: collagen bands which volume, density and crimp pattern are enhanced over time, more mature / controls</li></ul></td><td>[Young 1998]</td></tr></tbody></table>

<caption>(Table 4) Contd....</caption>
<table><thead><tr><th>Technique</th><th>Scaffold/parameters</th><th>Animal models / Sample size</th><th>Implantation site / Controls</th><th>Main results</th><th>Refs.</th></tr></thead><tbody><tr><td rowspan="2"></td><td>Bone marrow MSCs-collagen composites<br/>-Contraction: sutures</td><td>Rabbit<br/>2 groups (n=43 each):<br/>-with 1. 10<sup>6</sup>/ml cells and 4. 10<sup>6</sup>/mL cells composites in each tendon,<br/>-with 8.10<sup>6</sup>/mL cells in first tendon and defect left untreated in the second one<br/>8 animals to assess tendon normal properties</td><td>Patellar tendon central defects</td><td>At 12 and 26 weeks:<ul><li>Significant improvement of the biomechanical properties / natural repairs</li></ul>At 26 weeks:<ul><li>Maximum stress is 1/4 of normal tendon portion</li><li>No additional benefit of increasing cell density</li><li>Formation of ectopic bone in 28% of all composite grafted repairs</li></ul></td><td>[Awad 2003]</td></tr><tr><td>MSCs-collagen composites<br/>-Contraction: in silicon dishes between two posts</td><td>Rabbit<br/>2 groups :<br/>-with cells : 40.000 or 80.000 cells/mg collagen (n=16)<br/>- without cells (n=6)</td><td>Patellar tendon central defects</td><td>After 12 weeks:<ul><li>No significant difference in material and histological properties between the two cell concentrations and similar to acellular implants</li><li>Average linear modulus ~ 20% of normal values</li></ul></td><td>[Juncosa-Melvin 2005]</td></tr><tr><td rowspan="2">Plastic compression</td><td>Plastic-compressed collagen gels<br/>-Seeded or not with rabbit flexor tendon cells<br/>-20-40 µm thick<br/>-rolled to produce spiral rolls 22 mm in length x 2.25 mm in diameter</td><td>Rabbit<br/>Sacrificed at 3 time points:<br/>1 week (n=5), 3 weeks (n=4), 5 weeks (n=4)</td><td>Intercostal spaces<br/>3 cellular constructs on one side of the chest and 3 acellular constructs on the other one</td><td>After 5 weeks:<ul><li>Vascularization with a stronger response in the cellular constructs</li><li>Collagen remodeling and tensile mechanical strength enhanced in the cell-seeded gels</li></ul></td><td>[Mudera 2007]</td></tr><tr><td>Collagen type I sponge with rabbit bone marrow MSCs<br/>Either stimulated or non-stimulated constructs (2 weeks with 4% strain, 1 Hz cycle every 5 min, 8 h/day)</td><td>Rabbit<br/>n=10</td><td>Patellar tendon central defects</td><td>After 12 weeks:<ul><li>Maximal force 70% of normal tendons, vs 55% for non-stimulated constructs</li></ul></td><td>[Juncosa-Melvin 2006]<br/>[Juncosa-Melvin 2007]</td></tr><tr><td rowspan="2">Mechanical stimulation</td><td>Mixture of rabbit bone marrow MSCs and collagen gels combined with collagen sponges<br/>Either non stimulated or stimulated constructs (2 w. with 4% strain, 1 Hz cycle every 5 min, 8 h/day)</td><td>Rabbit<br/>n=6</td><td>Patellar tendon central defects</td><td>After 12 weeks:<ul><li>Averaged linear stiffness 80% of normal values, vs 50% for non-stimulated constructs</li><li>Maximal force 65% vs 60% for non-stimulated constructs</li></ul></td><td>[Shearn 2007]</td></tr></tbody></table>

biomechanical properties twelve weeks after surgery, the maximal force being 70% of the one sustained by normal tendons, versus 55% with non-stimulated constructs [227]. The stimulated constructs also presented a 2.5 rise in linear stiffness *in vitro* prior to implantation. The stimulation protocol was further optimized, and peak strain appeared to be a more important parameter than cycle

number and cycle repetition to enhance *in vitro* stiffness, with an optimized value of 2.4% (Table 3) [229]. Constructs made of a mixture of cells and collagen gels combined with collagen sponges showed a different behavior, with no significant change of the biomechanical properties measured prior to implantation in the
stimulated constructs, the improvement related to preconditioning being only present in the repair tissues [230].

## CONCLUDING REMARKS AND PERSPECTIVES

In the present review, we have seen how current tendon repair methods are mainly based on sutures and grafts that have proven to be effective while still exhibiting intrinsic drawbacks. We have seen that, even though tendon structure is not completely understood, tendon regeneration based on bioengineered scaffolds is promising. Among the different routes available, we focused on collagen-based scaffolds, as collagen I stands out from other biomolecules as being the main structural component of tendon, as well as for its suitable biocompatibility. The diversity of techniques and the increasing know-how in scaffold design allows tackling new exciting aspects such as alignment, multi-scale organization, scaffold tailoring or multi-cell deposition.

However, mechanical properties of collagen scaffolds still remain an issue while some other artificial materials compete or even overtake tendons. Still, several authors report that, while cross-linking improves mechanical properties and resistance to degradation, it may also damage collagen or decrease its biocompatibility. A fine balance must then be found. Cross-linking methods can be classified into two categories: chemical and physical cross-linking. Among chemical cross-linkers, widespread glutaraldehyde was proven to be cytotoxic [116]. EDC, that exhibits significant increase in mechanical properties may also decrease biocompatibility and collagen I production by cells [120]. More biological cross-linking methods such as lysyl-oxydation or glycation may be used [116]. Physical cross-linking can be achieved thanks to UV irradiation or dehydrothermal process. In some preclinical studies, the moderate mechanical properties of uncrosslinked collagen scaffolds have been shown to increase *in vivo*. Table 4 compares results obtained in animal models with various types of collagen scaffolds, among them cross-linked, compacted through cell-mediated contraction process, plastic-compressed and mechanically stimulated prior to implantation. After several weeks or months *in vivo*, most scaffolds exhibited partial replacement by neo-fibers and neo-vascularization in certain cases. The importance of mechanical pre-stimulation of cellularized scaffolds prior to implantation is highlighted in the works from Juncosa-Melvin *et al.* (2006, 2007) [227], [228], and Shearn *et al.* (2007) [230]. Twelve weeks after implantation in rabbit patellar tendon defects the maximal force obtained could reach 70% of normal tendons, which is comparable to the study led by Cavallaro *et al.* (1993) [118] using knitted EDC-crosslinked fibers with results obtained twelve weeks after implantation in dog anterior cruciate ligament.

Finally, this review shows that as bioengineering techniques improve, more complex tissues are achievable. Thus, some examples of myo-tendinous or bone-tendon junction reconstructions have been presented, illustrating the further increase of complexity. The development of artificial junctions could be paramount to improve overall scaffold efficiency and reliability, as well as to increase our understanding on the interactions between different biomaterials and cells.

## CONSENT FOR PUBLICATION

Not applicable.

## CONFLICT OF INTEREST

The authors declare no conflict of interest, financial or otherwise.

## ACKNOWLEDGEMENTS

The authors would like to thank David Pinto, Francisco M. Fernandes and Léa-Laetitia Pontani for critical reading of the manuscript. C. R. and L. P. were supported by a French Ministère de la Recherche fellowship.

## REFERENCES

[1] Morais DS, Torres J, Guedes RM, et al. Current approaches and future trends to promote tendon repair. Ann Biomed Eng 2015; 43: 2025-35.

[2] Thomopoulos S, Parks WC, Rifkin DB, et al. Mechanisms of tendon injury and repair: TENDON INJURY AND REPAIR. J Orthop Res 2015; 33: 832-9.

[3] Young JS, Maffulli N. Etiology and epidemiology of Achilles tendon problems. In: Maffulli N, Almekinders L, Eds. The Achilles Tendon. Springer 2007: pp. 39-49.

[4] Riley G. Tendinopathy—from basic science to treatment. Nat Clin Pract Rheumatol 2008; 4: 82-9.

[5] Lomas AJ, Ryan CNM, Sorushanova A, et al. The past, present and future in scaffold-based tendon treatments. Adv Drug Deliv Rev 2015; 84: 257-77.

[6] Lewis T, Cook J. Fluoroquinolones and Tendinopathy: A Guide for Athletes and Sports Clinicians and a Systematic Review of the Literature. J Athl Train 2014; 49: 422-27.

[7] Oliva F, Via AG, Maffulli N. Calcific tendinopathy of the rotator cuff tendons. Sports Med Arthrosc Rev 2011; 19: 237-43.

[8] Clayton RAE, Court-Brown CM. The epidemiology of musculoskeletal tendinous and ligamentous injuries. Injury 2008; 39: 1338-44.

[9] De Jong JP, Nguyen JT, Sonnema AJM, et al. The incidence of acute traumatic tendon injuries in the hand and wrist: A 10-Year Population-based Study. Clin Orthop Surg 2014; 6: 196.

[10] Sun C, Zhuo Q, Chai W, et al. Conservative interventions for treating Achilles tendon ruptures (Protocol). Cochrane Database of Systematic Reviews 2013. The Cochrane Collaboration. Published by John Wiley & Sons, Ltd. DOI: 10.1002/14651858.CD010765

[11] Scott A, Ashe MC. Common Tendinopathies in the Upper and Lower Extremities. Curr Sports Med Rep 2006; 5: 233-41.

[12] Sanders TL, Maradit KH, Bryan AJ, et al. The Epidemiology and Health Care Burden of Tennis Elbow: A Population-Based Study. Am J Sports Med 2015; 43: 1066-71.

[13] Pluim BM. Tennis injuries: occurrence, aetiology, and prevention. Br J Sports Med 2006; 40: 415-23.

[14] European Agency for Safety and Health at Work. OSH in figures: Work-related musculoskeletal disorders in the EU - Facts and figures; 2010. Available from: osha.europa.eu/fr/tools-and-publications/publications/reports/TERO09009ENC

[15] Health and Safety Executive, UK. Work-related Musculoskeletal disorder (WRMSDs) Statistics, Great Britain, 2015. Available from: hse.gov.uk/statistics/

[16] Agency for Healthcare Research and Quality. Comparative Effectiveness of the Nonoperative and Operative Treatments for Rotator Cuff Tears; 2010. Available from: ahrq.gov

[17] European Agency for Safety and Health at Work. Work-related neck and upper limb musculoskeletal disorders; 1999. Available from: osha.europa.eu/en/tools-and-publications/publications/reports/201

[18] Andarawis-Puri N, Flatow EL, Soslowsky LJ. Tendon basic science: Development, repair, regeneration, and healing. J Orthop Res 2015; 33: 780-4.

[19] Rawson S, Cartmell S, Wong J. Suture techniques for tendon repair; a comparative review. Muscles Ligaments Tendons J 2013; 3: 220.

[20] Squier CA, Bausch WH. Three-dimensional organization of fibroblasts and collagen fibrils in rat tail tendon. Cell Tissue Res 1984; 238: 319-27.

[21] Zhang G, Young BB, Ezura Y, et al. Development of tendon structure and function: regulation of collagen fibrillogenesis. J Musculoskelet Neuronal Interact 2005; 5: 5-21.

[22] Screen HRC, Berk DE, Kadler KE, et al. Tendon Functional Extracellular Matrix. J Orthop Res 2015; 33: 793-9.

[23] Kadler KE, Baldock C, Bella J, et al. Collagens at a glance. J Cell Sci 2007; 120: 1955-8.

[24] Pohlhammer J, O'Brien Jr WD, Dependence of the ultrasonic scatter coefficient on collagen concentration in mammalian tissues. J Acoust Soc Am 1981; 69: 283.

[25] Scott JE, Chemical morphology: The chemistry of our shape, in vivo and in vitro. Struct Chem 2007; 18: 257-65.

[26] Birch HL. Tendon matrix composition and turnover in relation to functional requirements. Int J Exp Pathol 2007; 88: 241-8.
[27] Bancelin S, Couture CA, Légaré K *et al.* Fast interferometric second harmonic generation microscopy. Biomed. Opt. Express 2016; 7: 399-408.

[28] Gusachenko I, Tran V, Houssen YG, et al. Polarization-Resolved Second-Harmonic Generation in Tendon upon Mechanical Stretching. Biophys J 2012; 102: 2220-9.

[29] Fouré A. New imaging methods for non-invasive assessment of mechanical, structural, and biochemical properties of human achilles tendon: A Mini Review. Front Physiol 2016; 7: 324.

[30] Hulmes DJS. Building Collagen Molecules, Fibrils, and Suprafirmillar Structures. J Struct Biol 2002; 137: 2-10.

[31] Benjamin M, Kaiser E, Milz S. Structure-function relationships in tendons: a review. J Anat 2008; 212: 211-28.

[32] Kannus P. Structure of the tendon connective tissue. Scand J Med Sci Sports 2000; 10: 312-20.

[33] Nicholls SP, Gathercole LJ, Keller A, et al. Crimping in rat tail tendon collagen: morphology and transverse mechanical anisotropy. Int J Biol Macromol 1983; 5: 283-8.

[34] Dlugosz J, Gathercole LJ, Keller A. Transmission electron microscope studies and their relation to polarizing optical microscopy in rat tail tendon. Micron 1978; 9: 71-82.

[35] Wang J H-C. Mechanobiology of tendon. J. Biomech 2006; 39: 1563-82.

[36] Shah JS, Palacios E, Palacios L. Development of crimp morphology and cellular changes in chick tendons. Dev Biol 1982; 94: 499-504.

[37] Herchenhan A, Kalson NS, Holmes DF, et al. Tenocyte contraction induces crimp formation in tendon-like tissue. Biomech Model Mechanobiol 2012; 11: 449-59.

[38] Hu XW, Knight DP, Chapman JA. The effect of non-polar liquids and non-ionic detergents on the ultrastructure and assembly of rat tail tendon collagen fibrils in vitro. Biochim Biophys Acta 1997; 1334: 327-37.

[39] Cartmell JS, Dunn MG. Effect of chemical treatments on tendon cellularity and mechanical properties. J. Biomed Mater Res 2000; 49: 134-40.

[40] Vidal B de C. Image analysis of tendon helical superstructure using interference and polarized light microscopy. Micron 2003; 34: 423-32.

[41] Grytz R, Meschke G. Constitutive modeling of crimped collagen fibrils in soft tissues. J Mech Behav Biomed Mater 2009; 2: 522-33.

[42] Reese SP, Maas SA, Weiss JA. Micromechanical models of helical superstructures in ligament and tendon fibers predict large Poisson's ratios. J Biomech 2010; 43: 1394-400.

[43] Livolant F, Leforestier A. Condensed phases of DNA: structures and phase transitions. Prog Polym Sci 1996; 21: 1115-64.

[44] Murthy NS. Liquid crystallinity in collagen solutions and magnetic orientation of collagen fibrils. Biopolymers 1984; 23: 1261-7.

[45] Maeda H. An atomic force microscopy study of ordered molecular assemblies and concentric ring patterns from evaporating droplets of collagen solutions. Langmuir 1999; 15: 8505-13.

[46] Giraud-Guille MM. Liquid crystallinity in condensed type I collagen solutions: a clue to the packing of collagen in extracellular matrices. J Mol Biol 1992; 224: 861-73.

[47] De Sa Peixoto P, Deniset-Besseau A, Schanne-Klein MC, et al. Quantitative assessment of collagen I liquid crystal organizations: role of ionic force and acidic solvent, and evidence of new phases. Soft Matter 2011; 7: 11203.

[48] Williams IF, Craig AS, Parry DAD, et al. Development of collagen fibril organization and collagen crimp patterns during tendon healing. Int J Biol Macromol 1985; 7: 275-82.

[49] Müller SA, Todorov A, Heisterbach PE et al. Tendon healing: an overview of physiology, biology, and pathology of tendon healing and systematic review of state of the art in tendon bioengineering. Knee Surg Sports Traumatol Arthrosc 2015; 23: 2097-105.

[50] Nourissat G, Berenbaum F, Duprez D. Tendon injury: from biology to tendon repair. Nat Rev Rheumatol 2015; 11: 223-33.

[51] Wada A, Kubota H, Miyanishi K, et al. Comparison of postoperative early active mobilization and immobilization in vivo utilising a four-strand flexor tendon repair. J Hand Surg Edinb Scotl 2001; 26: 301-6.

[52] Childress MA, Beutler A. Management of chronic tendon injuries. Am Fam Physician 2013; 87: 486-90.

[53] Khan KM, Scott A. Mechanotherapy: how physical therapists' prescription of exercise promotes tissue repair. Br J Sports Med 2009; 43: 247-52.

[54] Sundman EA, Cole BJ, Fortier LA. Growth factor and catabolic cytokine concentrations are influenced by the cellular composition of platelet-rich plasma. Am J Sports Med 2011; 39: 2135-40.

[55] Gelberman RH, Boyer MI, Brodt MD, et al. The effect of gap formation at the repair site on the strength and excursion of intrasynovial flexor tendons. An experimental study on the early stages of tendon-healing in dogs. J Bone Joint Surg Am 1999; 81: 975-82.

[56] Buckley PS, Pedowitz DI. Bone reduction clamp to gain length in repairing chronic achilles tendon ruptures. Orthopedics 2016; 39: e1223-5.

[57] Feldbrin Z, Hendel D, Lipkin A, et al. Achilles tendon rupture and our experience with the achillon device. Isr Med Assoc J 2010; 12: 609-12.

[58] Huffard B, O'Loughlin PF, Wright T et al. Achilles tendon repair: Achillon system vs. Krackow suture: An anatomic in vitro biomechanical study. Clin Biomech 2008; 23: 1158-64.

[59] Wang JHC. Mechanobiology of tendon. J Biomech 2006; 39: 1563-82.

[60] Gomes ME, Reis RL, Rodrigues MT, Eds. Tendon Regeneration: Understanding Tissue Physiology and Development to Engineer Functional Substitutes. Academic Press 2015.

[61] Maquirriain J. Achilles tendon rupture: Avoiding tendon lengthening during surgical repair and rehabilitation. Yale J Biol Med 2011; 84: 289-300.

[62] Schliemann B, Grüneweller N, Yao D, et al. Biomechanical evaluation of different surgical techniques for treating patellar tendon ruptures. Int Orthop 2016; 40: 1717-23.

[63] Hohendorff B, Siepen W, Spiering L, et al. Long-term results after operatively treated achilles tendon rupture: Fibrin glue versus suture. J Foot Ankle Surg 2008; 47: 392-9.

[64] Rose S, Prevoteau A, Elzière P, et al. Nanoparticle solutions as adhesives for gels and biological tissues. Nature 2014; 505: 382-5.

[65] Erol B, Kocaoğlu B, Esemenli T. Spiral-shaped metallic implant in the treatment of achilles tendon ruptures: An experimental study on the achilles tendon of sheep. J Foot Ankle Surg 2007; 46: 155-16.

[66] Ağır İ, Aytekin MN, Başçı O, et al. Tendon-holding capacities of two newly designed implants for tendon repair: An experimental study on the flexor digitorum profundus tendon of sheep. Open Orthop J 2014; 8: 135-9.

[67] Wolfe SW, Willis AA, Campbell D, et al. Biomechanic Comparison of the Teno Fix Tendon Repair Device With the Cruciate and Modified Kessler Techniques. J Hand Surg 2007; 32: 356-66.

[68] Lacoste S, Féron JM, Cherrier B. Percutaneous Tenolig® repair under intra-operative ultrasonography guidance in acute Achilles tendon rupture. Orthop Traumatol Surg Res 2014; 100: 925-30.

[69] Magnussen RA, Glisson RR, Moorman CT. Augmentation of achilles tendon repair with extracellular matrix xenograft a biomechanical analysis. Am J Sports Med 2011; 39: 1522-27.

[70] Beskin JL, Sanders RA, Hunter SC, et al. Surgical repair of Achilles tendon ruptures. Am J Sports Med 1987; 15: 1-8.

[71] Carmont MR, Maffulli N. Less invasive Achilles tendon reconstruction. BMC Musculoskelet Disord 2007; 8: 100.

[72] Ebed AM, Foda AAA. Less invasive reconstruction of chronic achilles tendon rupture with free semitendinosus tendon autograft - A case series. Int J Orthop 2016; 3: 539-43.

[73] Shewy MTE, Barbary HME, Abdel-Ghani H. Repair of chronic rupture of the achilles tendon using 2 intratendinous flaps from the proximal gastrocnemius-soleus complex. Am J Sports Med 2009; 37: 1570-7.

[74] Phipatanakul WP, Petersen SA. Porcine small intestine submucosa xenograft augmentation in repair of massive rotator cuff tears. Am J Orthop Belle Mead NJ 2009; 38: 572-5.

[75] Sclamberg SG, Tibone JE, Itamura JM, et al. Six-month magnetic resonance imaging follow-up of large and massive rotator cuff repairs reinforced with porcine small intestinal submucosa. J Shoulder Elb Surg 2004; 13: 538-41.

[76] Berlet GC, Hyer CF, Lee TH, et al. Collagen ribbon augmentation of achilles tendon tears: A biomechanical evaluation. J Foot Ankle Surg 2014; 53: 298-302.

[77] Song L, Olsen RE, Spalazzi JP, et al. Biomechanical evaluation of acellular collagen matrix augmented achilles tendon repair in sheep. J Foot Ankle Surg 2010; 49: 438-41.
[78] Lee DK. A preliminary study on the effects of acellular tissue graft augmentation in acute achilles tendon ruptures. J Foot Ankle Surg 2008; 47: 8-12.

[79] Hollawell S, Baione W. Chronic achilles tendon rupture reconstructed with achilles tendon allograft and xenograft combination. J Foot Ankle Surg 2015; 54: 1146-50.

[80] Herbert A, Jones GL, Ingham E. A biomechanical characterisation of acellular porcine super flexor tendons for use in anterior cruciate ligament replacement: Investigation into the effects of fat reduction and bioburden reduction bioprocesses. J Biomech 2015; 48: 22-9.

[81] Branch JP. A tendon graft weave using an acellular dermal matrix for repair of the achilles tendon and other foot and ankle tendons. J Foot Ankle Surg 2011; 50: 257-65.

[82] Gaspar D, Spanoudes K, Holladay C, et al. Progress in cell-based therapies for tendon repair. Adv Drug Deliv Rev 2015; 84: 240-56.

[83] Güngörüş C, Kolankaya D, and Aydin E. Histopathological and biomechanical evaluation of tenocyte seeded allografts on rat Achilles tendon regeneration. Biomaterials vol. 2015; 51: 108-18.

[84] Kew SJ, Gwynne JH, Enea D, et al. Regeneration and repair of tendon and ligament tissue using collagen fibre biomaterials. Acta Biomater 2011; 7: 3237-47.

[85] Stapp MD. Implantable materials and grafts in tendon surgery. In: Reconstructive surgery of the foot and leg: update '93. Tucker, GA: Podiatry Institute Publishing Co. 1993; pp. 289-92.

[86] Williams RD, August SF. Experimental evaluation of a Teflon tendon prosthesis. Am J Surg 1964; 107: 913-6.

[87] Holtz M, Midenberg ML, Kirschenbaum SE, Utilization of a silastic sheet in tendon repair of the foot. J Foot Surg 1982; 21: 253-9.

[88] Morrey MC, Barlow JD, Abdel MP, et al. Synthetic Mesh Augmentation of Acute and Subacute Quadriceps Tendon Repair. Orthopedics 2016; 39: e9-e13.

[89] Abdullah S. Usage of synthetic tendons in tendon reconstruction. BMC Proc 2015; 9: p. A68.

[90] Melvin A, Litsky A, Mayerson J, et al. An artificial tendon with durable muscle interface. J Orthop Res Off Publ Orthop Res Soc 2010; 28: 218-24.

[91] Jenkins DH, McKibbin B. The role of flexible carbon-fibre implants as tendon and ligament substitutes in clinical practice. A preliminary report. J Bone Joint Surg Br 1980; 62-B: 497-9.

[92] Corey SV. Tendon grafts and implants. In: Reconstructive surgery of the foot and leg: update '88. Tucker, GA: Podiatry Institute Publishing Co. 1988; pp. 58-61.

[93] Roberts JM, Goldstrohm GL, Brown TD, et al. Comparison of unrepaired, primarily repaired, and polyglactin mesh-reinforced Achilles tendon lacerations in rabbits. Clin Orthop 1983; 181: 244-9.

[94] Howard CB, McKibbin B, Ralis ZA. The use of Dexon as a replacement for the calcaneal tendon in sheep. Bone Jt J 1985; 67-B: 313-6.

[95] Liem MD, Zegel HG, Balduini FC, et al. Repair of Achilles tendon ruptures with a polylactic acid implant: assessment with MR imaging. AJR Am J Roentgenol 1991; 156: 769-73.

[96] Alexander H, Weiss AB, Parsons JR. Absorbable polymer-filamentous carbon composites--a new class of tissue scaffolding materials. Aktuelle Probl Chir Orthop 1983; 26: 78-91.

[97] Cao Y, Liu Y, Liu W, et al. Bridging tendon defects using autologous tenocyte engineered tendon in a hen model. Plast Reconstr Surg 2002; 110: 1280-9.

[98] Shoaib A, Mishra V. Surgical repair of symptomatic chronic achilles tendon rupture using synthetic graft augmentation. Foot Ankle Surg 2016. doi.org/10.1016/j.fas.2016.04.006

[99] Verdiyeva G, Koshy K, Glibbery N. Tendon reconstruction with tissue engineering approach—A review. J Biomed Nanotechnol 2015; 11: 1495-523.

[100] Ladd MR, Lee SJ, Stitzel JD, et al. Co-electrospun dual scaffolding system with potential for muscle-tendon junction tissue engineering. Biomaterials 2011; 32: 1549-59.

[101] Jung H-J, Fisher MB, Woo S L-Y. Role of biomechanics in the understanding of normal, injured, and healing ligaments and tendons. BMC Sports Sci Med Rehabil 2009; 1: 1-9.

[102] Tong WY, Shen W, Yeung CWF, et al. Functional replication of the tendon tissue microenvironment by a bioimprinted substrate and the support of tenocytic differentiation of mesenchymal stem cells. Biomaterials 2012; 33: 7686-98.

[103] Park SA, Kim IA, Lee YJ, et al. Biological responses of ligament fibroblasts and gene expression profiling on micropatterned sili-

cone substrates subjected to mechanical stimuli. J Biosci Bioeng 2006; 102: 402-12.

[104] Wang JH, Jia F, Gilbert TW, et al. Cell orientation determines the alignment of cell-produced collagenous matrix. J Biomech 2003; 36: 97-102.

[105] van Delft FCMJM, van den Heuvel FC, Loesberg WA, et al. Manufacturing substrate nano-grooves for studying cell alignment and adhesion. Microelectron Eng 2008; 85: 1362-6.

[106] Mertz L. What Is Biocompatibility?: A New Definition Based on the Latest Technology. IEEE Pulse 2013; 4: 14-5.

[107] Use of International Standard ISO 10993-1, 'Biological evaluation of medical devices - Part 1: Evaluation and testing within a risk management process'. U.S. Department of Health and Human Services Food and Drug Administration Center for Devices and Radiological Health, Jun 2016.

[108] Hutmacher DW. Scaffolds in tissue engineering bone and cartilage. Biomaterials 2000; 21: 2529-43.

[109] Badylak SF, Tullius R, Kokini K, et al. The use of xenogeneic small intestinal submucosa as a biomaterial for Achille's tendon repair in a dog model. J Biomed Mater Res 1995; 29: 977-85.

[110] Gilbert TW, Stewart-Akers AM, Simmons-Byrd A, et al. Degrada-tion and remodeling of small intestinal submucosa in canine achil-les tendon repair. J Bone Jt Surg Am 2007; 89: 621-30.

[111] Kew SJ, Gwynne JH, Enea D, et al., Regeneration and repair of tendon and ligament tissue using collagen fibre biomaterials. Acta Biomater 2011; 7: 3237-47.

[112] Glowacki J, Mizuno S. Collagen scaffolds for tissue engineering. Biopolymers 2008; 89: 338-44.

[113] Barczyk M, Carracedo S, Gullberg D. Integrins, Cell Tissue Res 2010; 339: 269-80.

[114] Jokinen J. Integrin-mediated cell adhesion to type I collagen fibrils. J Biol Chem 2004; 279: 31956-63.

[115] Murphy SV, Skardal A, Atala A. Evaluation of hydrogels for bio-printing applications. J Biomed Mater Res A 2013; 101A: 272-84.

[116] Walters BD, Stegemann JP. Strategies for directing the structure and function of three-dimensional collagen biomaterials across length scales. Acta Biomater 2014; 10: 1488-501.

[117] Muller SA, Durselen L, Heisterbach P, et al. Effect of a simple collagen type i sponge for achilles tendon repair in a rat model. Am J Sports Med 2016; 44: 1998-2004.

[118] Cavallaro JF, Kemp PD, Kraus KH. Collagen fabrics as biomaterials. Biotechnol Bioeng vol. 1994; 43: 781-91.

[119] Truhlsen SM, Fitzpatrick J. The extruded collagen suture: tissue reaction and absorption. Arch Ophthalmol 1965; 74: 371-4.

[120] Enea D, Henson F, Kew S, et al. Extruded collagen fibres for tissue engineering applications: effect of crosslinking method on mechanical and biological properties. J Mater Sci Mater Med 2011; 22: 1569-78.

[121] Zeugolis DI, Paul RG, Attenburrow G. Factors influencing the properties of reconstituted collagen fibers prior to self-assembly: Animal species and collagen extraction method. J Biomed Mater Res A 2008; 86A: 892-904.

[122] Pins GD, Christiansen DL, Patel R, et al. Self-assembly of collagen fibers. Influence of fibrillar alignment and decorin on mechanical properties. Biophys J 1997; 73: 2164-72.

[123] Wang MC, Pins GD, Silver FH. Collagen fibres with improved strength for the repair of soft tissue injuries. Biomaterials 1994; 15: 507-12.

[124] Lai ES, Anderson CM, Fuller GG. Designing a tubular matrix of oriented collagen fibrils for tissue engineering. Acta Biomater 2011; 7: 2448-56.

[125] Dunn MG, Avasarala PN, Zawadsky JP. Optimization of extruded collagen fibers for ACL reconstruction. J Biomed Mater Res 1993; 27: 1545-52.

[126] Kato YP, Christiansen DL, Hahn RA, et al. Mechanical properties of collagen fibres: a comparison of reconstituted and rat tail tendon fibres. Biomaterials 1989; 10: 38-42.

[127] Cornwell K. Collagen and Fibrin Biopolymer Microthreads for Bioengineered Ligament Generation: a Dissertation. GSBS Diss. Theses, May 2007.

[128] Huang Z-M, Zhang YZ, Kotaki, M, et al. A review on polymer nanofibers by electrospinning and their applications in nanocomposites. Compos Sci Technol 2003; 63: 2223-53.

[129] Teo WE, Ramakrishna S. A review on electrospinning design and nanofibre assemblies. Nanotechnology 2006; 17: R89-R106.
[130] Dong B, Arnoult O, Smith ME, et al. Electrospinning of Collagen Nanofiber Scaffolds from Benign Solvents. Macromol Rapid Commun 2009; 30: 539-42.

[131] Choktaweesap N, Arayanarakul K, Aht-Ong D, et al. Electrospun gelatin fibers: effect of solvent system on morphology and fiber diameters. Polym J 2007; 39: 622-31.

[132] Wan YQ, He JH, Yu JY, et al. Electrospinning of high-molecule PEO solution. J Appl Polym Sci 2007; 103: 3840-3.

[133] Bürck J, Heissler, Geckler U, et al. Resemblance of electrospun collagen nanofibers to their native structure. Langmuir 2013; 29: 1562-72.

[134] Matthews JA, Wnek GE, Simpson DG, et al. Electrospinning of collagen nanofibers. Biomacromolecules 2002; 3: 232-8.

[135] Matthews JA, Boland ED, Wnek GE, et al. Electrospinning of collagen type II: a feasibility study. J Bioact Compat Polym 2003; 18: 125-34.

[136] Rho KS, Jeong L, Lee G, et al. Electrospinning of collagen nanofibers: Effects on the behavior of normal human keratinocytes and early-stage wound healing. Biomaterials, 2006; 27: 1452-61.

[137] Zhong SP, Teo WE, Zhu X, et al. Development of a novel collagen-GAG nanofibrous scaffold via electrospinning. Mater Sci Eng C 2007; 27: 262-6.

[138] Zeugolis DI, Khew ST, Yew ESY, et al. Electro-spinning of pure collagen nano-fibres - Just an expensive way to make gelatin? Biomaterials 2008; 29: 2293-305.

[139] Szentivanyi A, Assmann U, Schuster R, et al. Production of biohybrid protein/PEO scaffolds by electrospinning. Mater Werkst 2009; 40: 65-72.

[140] Buttafoco L, Kolkman NG, Engbers-Buijtenhuijs P, et al. Electro-spinning of collagen and elastin for tissue engineering applications. *Biomaterials* 2006; 27: 724-34.

[141] Elamparithi A, Punnoose AM, Kuruvilla S. Electrospun type 1 collagen matrices preserving native ultrastructure using benign binary solvent for cardiac tissue engineering. Artif Cells Nanomedicine Biotechnol 2016; 44: 1318-25.

[142] Xu Y, Wu J, Wang H, et al. Fabrication of electrospun poly(L-Lactide-co-ε-Caprolactone)/collagen nanoyarn network as a novel, three-dimensional, macroporous, aligned scaffold for tendon tissue engineering. Tissue Eng Part C Methods 2013; 19: pp. 925-36.

[143] Deville S. Ice-templating, freeze casting: Beyond materials processing. J Mater Res 2013; 28: 2202-19.

[144] Haugh MG, Murphy CM, O'Brien FJ. Novel freeze-drying methods to produce a range of collagen-glycosaminoglycan scaffolds with tailored mean pore sizes. Tissue Eng Part C Methods 2009; 16: 887-94.

[145] Pawelec KM, Husmann A, Best SM, et al. A design protocol for tailoring ice-templated scaffold structure. J R Soc Interface 2014; 11: 1.

[146] Pawelec KM, Husmann A, Best SM, et al. Understanding anisotropy and architecture in ice-templated biopolymer scaffolds. Mater Sci Eng C 2014; 37: 141-7.

[147] Pawelec KM, Husmann A, Wardale RJ, et al. Ionic solutes impact collagen scaffold bioactivity. J Mater Sci Mater Med 2015; 26: 91.

[148] Pawelec KM, Husmann A, Best SM, et al. Altering crystal growth and annealing in ice-templated scaffolds. J Mater Sci 2015; 50: 7537-43.

[149] Pawelec KM, Wardale RJ, Best SM, et al. The effects of scaffold architecture and fibrin gel addition on tendon cell phenotype. J Mater Sci Mater Med 2015; 5349.

[150] Davidenko N, Bax DV, Schuster CF et al. Optimisation of UV irradiation as a binding site conserving method for crosslinking collagen-based scaffolds. J Mater Sci Mater Med 2016; 27: 14.

[151] Lowe CJ, Reucroft IM, Grota MC, et al. Production of highly aligned collagen scaffolds by freeze-drying of self-assembled, fibrillar collagen gels. ACS Biomater Sci Eng 2016; 2: 643-51.

[152] Caliari SR, Harley BAC. The effect of anisotropic collagen-GAG scaffolds and growth factor supplementation on tendon cell recruitment, alignment, and metabolic activity. Biomaterials 2011; 32: 5330-40.

[153] Caliari SR, Harley BAC. Composite Growth Factor Supplementation Strategies to Enhance Tenocyte Bioactivity in Aligned Collagen-GAG Scaffolds. Tissue Eng Part A 2013; 19: 1100-12.

[154] Caliari SR, Ramirez MA, Harley BAC. The development of collagen-GAG scaffold-membrane composites for tendon tissue engineering. Biomaterials 2011; 32: 8990-8.

[155] Caliari SR, Weisgerber DW, Grier WK, et al. Collagen scaffolds incorporating coincident gradations of instructive structural and biochemical cues for osteotendinous junction engineering. Adv Healthc Mater 2015; 4: 831-7.

[156] Mozdzen LC, Rodgers R, Banks JM, et al. Increasing the strength and bioactivity of collagen scaffolds using customizable arrays of 3D-printed polymer fibers. Acta Biomater 2016; 33: 25-33.

[157] McCullough EJ, Yadavalli VK. Surface modification of fused deposition modeling ABS to enable rapid prototyping of biomedical microdevices. J Mater Process Technol 2013; 213: 947-54.

[158] Murphy SV, AtalaA. 3D bioprinting of tissues and organs. Nat Biotechnol 2014; 32: 773-85.

[159] Derby B. Printing and Prototyping of Tissues and Scaffolds. Science 2012; 338: 921-6.

[160] Do AV, Khorsand B, Geary SM, et al. 3D printing of scaffolds for tissue regeneration applications. Adv Healthc Mater 2015; 4: 1742-62.

[161] Inzana JA, Olvera D, Fuller SM, et al. 3D printing of composite calcium phosphate and collagen scaffolds for bone regeneration. Biomaterials 2014; 35: 4026-34.

[162] Duarte Campos DF, Blaeser A, Buellesbach K, et al. Bioprinting organotypic hydrogels with improved mesenchymal stem cell remodeling and mineralization properties for bone tissue engineering. Adv Healthc Mater 2016; 5: 1336-45.

[163] Lin K-F, He S, Song Y, et al. Low-temperature additive manufacturing of biomimic three-dimensional hydroxyapatite/collagen scaffolds for bone regeneration. ACS Appl Mater Interfaces, 2016; 8: 6905-16.

[164] Park JY, Choi JC, Shim J-H, et al. A comparative study on collagen type I and hyaluronic acid dependent cell behavior for osteochondral tissue bioprinting. Biofabrication 2014; 6: 035004.

[165] Legemate K, Tarafder S, Jun Y, et al. Engineering Human TMJ Discs with Protein-Releasing 3D-Printed Scaffolds. J Dent Res 2016; 95: 800-7.

[166] Lee HJ, Kim YB, Ahn SH, et al. A New Approach for Fabricating Collagen/ECM-Based Bioinks Using Preosteoblasts and Human Adipose Stem Cells. Adv Healthc Mater 2015; 4: 1359-68.

[167] Wu Z, Su X, Xu Y, et al. Bioprinting three-dimensional cell-laden tissue constructs with controllable degradation. Sci Rep 2016; 6: 24474.

[168] Xu T, Binder KW, Albanna M, et al. Hybrid printing of mechanically and biologically improved constructs for cartilage tissue engineering applications. Biofabrication 2013; 5: 015001.

[169] Sun K, Li R, Jiang W, et al. Comparison of three-dimensional printing and vacuum freeze-dried techniques for fabricating composite scaffolds. Biochem Biophys Res Commun 2016; 477: 1085-91.

[170] Lode A, Meyer M, Brüggemeier S, et al. Additive manufacturing of collagen scaffolds by three-dimensional plotting of highly viscous dispersions. Biofabrication 2016; 8: 015015.

[171] Liu CZ, Xia ZD, Han ZW, et al. Novel 3D collagen scaffolds fabricated by indirect printing technique for tissue engineering. J Bio-med Mater Res B Appl Biomater 2008; 85: 519-28.

[172] Ahn S, Lee S, Cho Y, et al. Fabrication of three-dimensional collagen scaffold using an inverse mould-leaching process. Bioprocess Biosyst Eng 2011; 34: 903-11.

[173] Yeo M, Lee J-S, Chun W, et al. An innovative collagen-based cell-printing method for obtaining human adipose stem cell-laden structures consisting of core-sheath structures for tissue engineering. Biomacromolecules 2016; 17: 1365-75.

[174] Moon S, Hasan SK, Song YS, et al. Layer by layer three-dimensional tissue epitaxy by cell-laden hydrogel droplets. Tissue Eng Part C Methods 2009; 16: 157-66.

[175] Kang H-W, Lee SJ, Ko IK, et al. A 3D bioprinting system to produce human-scale tissue constructs with structural integrity. Nat Biotechnol 2016; 34: 312-9.

[176] Merceron TK, Burt M, Seol Y-J, et al. A 3D bioprinted complex structure for engineering the muscle-tendon unit. Biofabrication 2015; 7: 035003.

[177] Baker HR, Merschrod S EF, Poduska KM. Electrochemically Controlled Growth and Positioning of Suspended Collagen Membranes. Langmuir 2008; 24: 2970-2.

[178] Cheng X, Gurkan UA, Dehen CJ, et al. An electrochemical fabrication process for the assembly of anisotropically oriented collagen bundles. Biomaterials 2008; 29: 3278-88.
[179] Younesi M, Islam A, Kishore V, et al. Fabrication of composition-ally and topographically complex robust tissue forms by 3D-electrochemical compaction of collagen. Biofabrication 2015; 7: 035001.

[180] Abu-Rub MT, Billiar KL, van Es MH, et al. Nano-textured self-assembled aligned collagen hydrogels promote directional neurite guidance and overcome inhibition by myelin associated glycoprotein. Soft Matter 2011; 7: 2770-81.

[181] Uquillas JA, Kishore V, Akkus O, Effects of phosphate-buffered saline concentration and incubation time on the mechanical and structural properties of electrochemically aligned collagen threads. Biomed Mater 2011; 6: 035008.

[182] Kishore V, Bullock W, Sun X, et al. Tenogenic differentiation of human MSCs induced by the topography of electrochemically aligned collagen threads. Biomaterials 2012; 33: 2137-44.

[183] Kishore V, Uquillas JA, Dubikovsky A, et al. In vivo response to electrochemically aligned collagen bioscaffolds. J Biomed Mater Res B 2012; 100B: 400-8.

[184] Uquillas JA, Kishore V, Akkus O. Genipin crosslinking elevates the strength of electrochemically aligned collagen to the level of tendons. J Mech Behav Biomed Mater 2012; 15: 176-89.

[185] Younesi M, Islam A, Kishore V, et al. Tenogenic Induction of Human MSCs by Anisotropically Aligned Collagen Biotextiles. Adv Funct Mater 2014; 24: 5762-70.

[186] Islam A, Chapin K, Younesi M, et al. Computer aided biomanufacturing of mechanically robust pure collagen meshes with controlled macroporosity. Biofabrication 2015; 7: 035005.

[187] Bell E, Ivarsson B, Merrill C. Production of a tissue-like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential in vitro. Proc Natl Acad Sci 1979; 76: 1274-8.

[188] Stopak D, Harris AK. Connective tissue morphogenesis by fibroblast traction: I. Tissue culture observations. Dev Biol 1982; 90: 383-98.

[189] Barocas VH, Tranquillo RT. An anisotropic biphasic theory of tissue-equivalent mechanics: the interplay among cell traction, fibrillar network deformation, fibril alignment, and cell contact guidance. J Biomech Eng 1997; 119: 137-45.

[190] Huang D, Chang TR, Aggarwal A, et al. Mechanisms and dynamics of mechanical strengthening in ligament-equivalent fibroblast-populated collagen matrices. Ann Biomed Eng 1993; 21: 289-305.

[191] Young RG, Butler DL, Weber W, et al. Use of mesenchymal stem cells in a collagen matrix for Achilles tendon repair. J Orthop Res 1998; 16: 406-13.

[192] Awad HA, Boivin GP, Dressler MR, et al. Repair of patellar tendon injuries using a cell-collagen composite. J Orthop Res 2003; 21: 420-31.

[193] Butler DL, Juncosa-Melvin N, Boivin GP, et al. Functional tissue engineering for tendon repair: A multidisciplinary strategy using mesenchymal stem cells, bioscaffolds, and mechanical stimulation. J Orthop Res 2008; 26: 1-9.

[194] Juncosa-Melvin N, Boivin GP, Galloway MT, et al. Effects of cell-to-collagen ratio in mesenchymal stem cell-seeded implants on tendon repair biomechanics and histology. Tissue Eng 2005; 1: 448-57.

[195] Calve S, Dennis RG, Kosnik PE, et al. Engineering of functional tendon. Tissue Eng 2004; 10: 755-61.

[196] Swasdison S, Mayne R. Formation of highly organized skeletal muscle fibers in vitro. Comparison with muscle development in vivo. J Cell Sci 1992; 102: 643-52.

[197] Larkin LM, Calve S, Kostrominova TY, et al. Structure and Functional Evaluation of Tendon-Skeletal Muscle Constructs Engineered in Vitro. Tissue Eng 2006; 12: 3149-58.

[198] Kapacee Z, Richardson SH, Lu Y, et al., Tension is required for fibripositor formation. Matrix Biol 2008; 27: 371-5.

[199] Guerquin M-J, Charvet B, et al., Nourissat G, et al. Transcription factor EGR1 directs tendon differentiation and promotes tendon repair. J Clin Invest 2013; 123: 3564-76.

[200] Neal D, Sakar MS, Ong L-LS, et al. Formation of elongated fascicle-inspired 3D tissues consisting of high-density, aligned cells using sacrificial outer molding. Lab Chip 2014; 14: 1907.

[201] Yeung C-YC, Zeef LAH, Lallyett C, et al. Chick tendon fibroblast transcriptome and shape depend on whether the cell has made its own collagen matrix. Sci Rep 2015; 5: 13555.

[202] Breidenbach AP, Dyment NA, Lu Y, et al. Fibrin gels exhibit improved biological, structural, and mechanical properties compared

with collagen gels in cell-based tendon tissue-engineered constructs. Tissue Eng Part A 2015; 21: 438-50.

[203] Torbet J, Ronziere M-C. Magnetic alignment of collagen during self-assembly. Biochem J 1984; 219: 1057-9.

[204] Guido S, Tranquillo RT. A methodology for the systematic and quantitative study of cell contact guidance in oriented collagen gels. Correlation of fibroblast orientation and gel birefringence. J Cell Sci 1993; 105: 317-31.

[205] Chen S, Hirota N, Okuda M, et al. Microstructures and rheological properties of tilapia fish-scale collagen hydrogels with aligned fibrils fabricated under magnetic fields. Acta Biomater 2011; 7: 644-52.

[206] Guo C, Kaufman LJ. Flow and magnetic field induced collagen alignment. Biomaterials 2007; 28: 1105-14.

[207] Lee P, Lin R, Moon J, et al. Microfluidic alignment of collagen fibers for in vitro cell culture. Biomed Microdevices 2006; 8: 35-41.

[208] Saeidi N, Sander EA, Ruberti JW. Dynamic shear-influenced collagen self-assembly. Biomaterials 2009; 30: 6581-92.

[209] Lanfer B, Freudenberg U, Zimmermann R, et al. Aligned fibrillar collagen matrices obtained by shear flow deposition. Biomaterials 2008; 29: 3888-95.

[210] Haynl C, Hofmann E, Pawar K, et al. Microfluidics-produced collagen fibers show extraordinary mechanical properties. Nano Lett 2016; 16: 5917-22.

[211] Yunoki S, Hatayama H, Ebisawa M, et al. A novel fabrication method to create a thick collagen bundle composed of uniaxially aligned fibrils: An essential technology for the development of artificial tendon/ligament matrices. J Biomed Mater Res A 2015; 103: 3054-65.

[212] Paten JA, Siadat SM, Susilo ME, et al. Flow-Induced Crystallization of Collagen: A Potentially Critical Mechanism in Early Tissue Formation. ACS Nano 2016; 10: 5027-40.

[213] Brown RA, Wiseman M, Chuo C-B, et al. Ultrarapid engineering of biomimetic materials and tissues: fabrication of nano- and microstructures by plastic compression. Adv Funct Mater 2005; 15: 1762-70.

[214] Ghezzi CE, Muja N, Marelli B, et al. Real time responses of fibroblasts to plastically compressed fibrillar collagen hydrogels. Biomaterials 2011; 32: 4761-72.

[215] Mudera V, Morgan M, Cheema U, et al. Ultra-rapid engineered collagen constructs tested in an in vivo nursery site. J Tissue Eng Regen Med 2007; 1: 192-8.

[216] Marelli B, Ghezzi CE, James-Bhasin M, et al. Fabrication of injectable, cellular, anisotropic collagen tissue equivalents with modular fibrillar densities. Biomaterials 2015; 37: 183-93.

[217] Killian ML, Cavinatto L, Galatz LM, et al. The role of mechanobiology in tendon healing. J Shoulder Elbow Surg 2012; 21: 228-37.

[218] Pins GD, Huang EK, Christiansen DL, et al. Effect of static axial strain on the tensile properties and failure mechanisms of self-assembled collagen fibers. J Appl Polym Sci 1997; 63: 1429-40.

[219] Chaubaroux C, Perrin-Schmitt F, Senger B, et al. Cell alignment driven by mechanically induced collagen fiber alignment in collagen/alginate coatings. Tissue Eng Part C Methods 2015; 21: 881-8.

[220] Cheema U, Chuo C-B, Sarathchandra P, et al. Engineering functional collagen scaffolds: Cyclical loading increases material strength and fibril aggregation. Adv Funct Mater 2007; 17: 2426-31.

[221] Zeichen J, van Griensven M, Bosch U. The proliferative response of isolated human tendon fibroblasts to cyclic biaxial mechanical strain. Am J Sports Med 2000; 28: 888-92.

[222] Riboh J, Chong AKS, Pham H, et al. Optimization of flexor tendon tissue engineering with a cyclic strain bioreactor. J Hand Surg 2008; 33: 1388-96.

[223] Zhang L, Kahn CJF, Chen H-Q, et al. Effect of uniaxial stretching on rat bone mesenchymal stem cell: Orientation and expressions of collagen types I and III and tenascin-C. Cell Biol Int 2008; 32: 344-52.

[224] Garvin J, Qi J, Maloney M, et al. Novel system for engineering bioartificial tendons and application of mechanical load. Tissue Eng 2003; 9: 967-79.

[225] Kuo CK, Tuan RS. Mechanoactive Tenogenic Differentiation of Human Mesenchymal Stem Cells. Tissue Eng Part A 2008; 14: 1615-27.
[226] Scott A, Danielson P, Abraham T, et al. Mechanical force modulates scleraxis expression in bioartificial tendons. J Musculoskelet Neuronal Interact 2011; 11: 124-32.

[227] Juncosa-Melvin N, Shearn JT, Boivin GP et al. Effects of mechanical stimulation on the biomechanics and histology of stem cell-collagen sponge constructs for rabbit patellar tendon repair. Tissue Eng 2006; 12: 2291-300.

[228] Juncosa-Melvin N, Matlin KS, Holdcraft RW, et al. Mechanical Stimulation Increases Collagen Type I and Collagen Type III Gene

Expression of Stem Cell-Collagen Sponge Constructs for Patellar Tendon Repair. Tissue Eng 2007; 13: 1219-26.

[229] Nirmalanandhan VS, Shearn JT, Juncosa-Melvin N et al. Improving linear stiffness of the cell-seeded collagen sponge constructs by varying the components of the mechanical stimulus. Tissue Eng. Part A 2008; 14: 1883-91.

[230] Shearn JT, Juncosa-Melvin N, Boivin GP et al. Mechanical stimulation of tendon tissue engineered constructs: effects on construct stiffness, repair biomechanics, and their correlation. J Biomech Eng 2007; 129: 848-54.
List of Figures

<table><tr><td>1.1 Scheme of the tendon location</td><td>6</td></tr><tr><td>1.2 Scheme of the hierarchical organization of the tendon</td><td>7</td></tr><tr><td>1.3 Scheme of a equine superficial flexor tendon</td><td>8</td></tr><tr><td>1.4 Scheme of collagen fibers and fibrils organization</td><td>8</td></tr><tr><td>1.5 Schemes illustrating the major structural characteristics of extracellular matrix of tendon</td><td>9</td></tr><tr><td>1.6 Scheme conciliating helix crimp and linear organization</td><td>10</td></tr><tr><td>1.7 Scheme of collagen fibers and fibrils organization</td><td>11</td></tr><tr><td>1.8 Location, annual incidence and main cause of most frequent tendon injuries</td><td>14</td></tr><tr><td>1.9 Scheme of the three sutures techniques currently used</td><td>16</td></tr><tr><td>1.10 Images of different metallic implants for tendon repair</td><td>17</td></tr><tr><td>1.11 Scheme of Achilles tendon rupture repair by using autologous tendon</td><td>19</td></tr><tr><td>1.12 Images of commercially available A: OrthADAPT™ xenograft and B: Graf-jacket® allograft.</td><td>20</td></tr><tr><td>1.13 Image of commercially available synthetic graft: Artelon®.</td><td>22</td></tr><tr><td>1.14 Pictures of different metallic implants for tendon repair</td><td>23</td></tr><tr><td>1.15 Schematic of the techniques used to produce scaffolds for tendon repair</td><td>25</td></tr><tr><td>1.16 PGA/PLA scaffold example</td><td>26</td></tr><tr><td>1.17 Braided chitosan scaffolds</td><td>28</td></tr><tr><td>1.18 3D plotted collagen scaffold (33 layers) in dry state</td><td>31</td></tr><tr><td>2.1 Experimental set-up of extrusion scheme</td><td>47</td></tr><tr><td>2.2 Example of a complex fluid extrusion</td><td>48</td></tr><tr><td>2.3 Example of a 23G needle tip</td><td>49</td></tr><tr><td>2.4 Protocol of stainless steel blunt needles preparation</td><td>50</td></tr><tr><td>2.5 Protocol of glass needles fabrication</td><td>51</td></tr><tr><td>2.6 Images of fire-polished glass capillary</td><td>51</td></tr><tr><td>2.7 Piston stick-slip behaviors</td><td>52</td></tr><tr><td>2.8 Pistons</td><td>53</td></tr><tr><td>2.9 Scheme of the { syringe + capillary } system and the characteristic variables</td><td>53</td></tr><tr><td>2.10 Force signals and their respective piston velocities obtained for alginate solution extrusion</td><td>55</td></tr><tr><td>2.11 Image of extrusion with a glass capillary where air bubbles are entrapped in the solution</td><td>56</td></tr><tr><td>2.12 Scheme of a cylinder pipe of length L and radius R.</td><td>57</td></tr><tr><td>2.13 Wall shear stress as a function of shear rate for glass capillaries of different radii</td><td>58</td></tr><tr><td>2.14 Velocity profiles for the flow of a fluid obeying a power law.</td><td>59</td></tr></table>
<table>
  <tbody>
    <tr>
      <td>2.15 Velocity profile for a single shear flow of a newtonian liquid between two parallel plates.</td>
      <td>60</td>
    </tr>
    <tr>
      <td>2.16 Scheme of shear flow tensor definition in 3 dimensions</td>
      <td>61</td>
    </tr>
    <tr>
      <td>2.17 Maxwell liquid model scheme</td>
      <td>63</td>
    </tr>
    <tr>
      <td>2.18 Maxwell fluid behavior for an oscillatory shear</td>
      <td>64</td>
    </tr>
    <tr>
      <td>2.19 Rheometer geometries: left: Double gap Couette cell, middle: Parallel plates and right: Cone and plate</td>
      <td>67</td>
    </tr>
    <tr>
      <td>2.20 Two configurations of mechanical testing</td>
      <td>70</td>
    </tr>
    <tr>
      <td>2.21 Table of collagen families</td>
      <td>71</td>
    </tr>
    <tr>
      <td>2.22 Collagen biosynthesis scheme</td>
      <td>72</td>
    </tr>
    <tr>
      <td>2.23 Collagen type I triple helix scheme.</td>
      <td>73</td>
    </tr>
    <tr>
      <td>2.24 Collagen nat charge plot as a function of the pH</td>
      <td>74</td>
    </tr>
    <tr>
      <td>2.25 Rat tail tendons anatomy and extraction</td>
      <td>76</td>
    </tr>
    <tr>
      <td>2.26 Electrophoresis gel</td>
      <td>77</td>
    </tr>
    <tr>
      <td>2.27 Centrifugal concentrator scheme</td>
      <td>78</td>
    </tr>
    <tr>
      <td>2.28 Sodium alginate chemical structures</td>
      <td>79</td>
    </tr>
    <tr>
      <td>2.29 Alginate persistence length as a function of Ionic Strength and pH</td>
      <td>80</td>
    </tr>
    <tr>
      <td>2.30 Schematic representation of the "egg-box" structure</td>
      <td>81</td>
    </tr>
    <tr>
      <td>2.31 Schematic illustration of the multiple-step binding of Ca<sup>2+</sup> to alginate</td>
      <td>82</td>
    </tr>
    <tr>
      <td>2.32 Proteins and polysaccharides mixtures schematic representation</td>
      <td>83</td>
    </tr>
    <tr>
      <td>2.33 Solutions aspects before being extruded</td>
      <td>84</td>
    </tr>
    <tr>
      <td>3.1 Collagen Shear storage and loss moduli as a function of the angular frequency</td>
      <td>89</td>
    </tr>
    <tr>
      <td>3.2 Prefactors G'<sub>0</sub>, G''<sub>0</sub> and power-law exponent β as a function of the collagen concentration</td>
      <td>89</td>
    </tr>
    <tr>
      <td>3.3 Comparison of the complex (or dynamic) viscosity η* for a collagen solution at 30 mg/mL</td>
      <td>90</td>
    </tr>
    <tr>
      <td>3.4 Characteristic disentanglement time τ<sub>c</sub> as a function of the collagen concentration</td>
      <td>91</td>
    </tr>
    <tr>
      <td>3.5 Shear rate dependent Weissenberg numbers for collagen solution at 30 mg/mL.</td>
      <td>92</td>
    </tr>
    <tr>
      <td>3.6 Viscosity of collagen at different concentrations as a function of the shear rate.</td>
      <td>94</td>
    </tr>
    <tr>
      <td>3.7 Specific viscosity of alginate solutions as a function of concentration (wt %)</td>
      <td>95</td>
    </tr>
    <tr>
      <td>3.8 Shear-thinning behavior of the alginate solutions.</td>
      <td>96</td>
    </tr>
    <tr>
      <td>3.9 Alginate shear storage G' (empty circle) and loss G" (filled circle) moduli against angular frequencies</td>
      <td>97</td>
    </tr>
    <tr>
      <td>3.10 Alginate first normal stress difference measured in steady state shear (empty circles) and computed from Laun empirical relationship</td>
      <td>98</td>
    </tr>
    <tr>
      <td>3.11 Alginate shear storage G' (empty circle) and loss G" (filled circle) moduli against angular frequencies</td>
      <td>98</td>
    </tr>
    <tr>
      <td>4.1 Examples of polymer melt extrudates</td>
      <td>103</td>
    </tr>
    <tr>
      <td>4.2 Morphologies observed during the extrusion of alginate while changing the polymer concentration, and die material without binding ions</td>
      <td>106</td>
    </tr>
    <tr>
      <td>4.3 Morphologies observed during the extrusion of alginate while changing the polymer concentration, and die material without binding ions for higher velocities</td>
      <td>107</td>
    </tr>
    <tr>
      <td>4.4 Path of train of bubbles during the extrusion</td>
      <td>108</td>
    </tr>
    <tr>
      <td>4.5 Interlocking of two capillaries of radii 250 and 150 µm</td>
      <td>108</td>
    </tr>
    <tr>
      <td>4.6 Example of protocols used to produce wavy scaffolds.</td>
      <td>122</td>
    </tr>
  </tbody>
</table>
<table>
  <tbody>
    <tr>
      <td>4.7</td>
      <td>Collagen at 30 mg/mL extrusion in A) PBS 1X and B) PBS 5X observed in real time and post extrusion</td>
      <td>123</td>
    </tr>
    <tr>
      <td>4.8</td>
      <td>Morphologies observed during the extrusion of sodium alginate solution at 4% in CaCl<sub>2</sub>0.5M</td>
      <td>125</td>
    </tr>
    <tr>
      <td>4.9</td>
      <td>Ratio diameters D'/D plot as a function of the shear rate for the extrusion of ALG4 in CaCl<sub>2</sub>0.5M with stainless steel needles</td>
      <td>126</td>
    </tr>
    <tr>
      <td>4.10</td>
      <td>Steady and homogeneous alginate flow and the scheme representation of the syneresis process occuring</td>
      <td>128</td>
    </tr>
    <tr>
      <td>4.11</td>
      <td>Die swelling image and the respective plot of swelling ratio for stainless stell needles</td>
      <td>129</td>
    </tr>
    <tr>
      <td>4.12</td>
      <td>The Weissenberg or rod-climbing effect imaging</td>
      <td>129</td>
    </tr>
    <tr>
      <td>4.13</td>
      <td>Scheme of die swelling mechanism</td>
      <td>130</td>
    </tr>
    <tr>
      <td>4.14</td>
      <td>Extrusion of ALG4 in CaCl<sub>2</sub>0.5M (left) and CaCl<sub>2</sub>0.1M (right) with glass capillary R=250μm at the same velocity.</td>
      <td>131</td>
    </tr>
    <tr>
      <td>4.15</td>
      <td>Laser diffraction experiment on an alginate thread which exhibits periodic striations</td>
      <td>132</td>
    </tr>
    <tr>
      <td>4.16</td>
      <td>Eye guidelines of the possible mechanism which leads to helical (left) and striated pattern (right)</td>
      <td>132</td>
    </tr>
    <tr>
      <td>4.17</td>
      <td>Striations velocity evolution as a function of the shear rate for three capillaries of radii</td>
      <td>133</td>
    </tr>
    <tr>
      <td>4.18</td>
      <td>Striations velocity evolution as a function of the shear rate for three capillaries of radii</td>
      <td>134</td>
    </tr>
    <tr>
      <td>4.19</td>
      <td>Spatio-temporal image of ALG4 extrusion with stainless steel needle of radii 760μm in the oscillating regime.</td>
      <td>134</td>
    </tr>
    <tr>
      <td>4.20</td>
      <td>Gel clogging steps</td>
      <td>135</td>
    </tr>
    <tr>
      <td>4.21</td>
      <td>Spatio-temporal images of ALG4 extrusion</td>
      <td>136</td>
    </tr>
    <tr>
      <td>4.22</td>
      <td>A) Force and velocity signal of the extrusion of ALG4 in CaCl<sub>2</sub>0.5M with a glass capillary of 500μm.</td>
      <td>136</td>
    </tr>
    <tr>
      <td>5.1</td>
      <td>Scheme of a linearly polarized sinusoidal electromagnetic wave travelling in the positive x-direction.</td>
      <td>144</td>
    </tr>
    <tr>
      <td>5.2</td>
      <td>Index ellipsoid for a positive uniaxial medium</td>
      <td>145</td>
    </tr>
    <tr>
      <td>5.3</td>
      <td>Excitation of molecules with a radiation of frequency ω.</td>
      <td>147</td>
    </tr>
    <tr>
      <td>5.4</td>
      <td>Schemes of the origin of the collagen signal</td>
      <td>148</td>
    </tr>
    <tr>
      <td>5.5</td>
      <td>Scheme of SHG microscope set-up and the main orientation parameters definition</td>
      <td>149</td>
    </tr>
    <tr>
      <td>5.6</td>
      <td>The two angles φ and ψ describe the orientation of the collagen molecules in the excitation volume</td>
      <td>150</td>
    </tr>
    <tr>
      <td>5.7</td>
      <td>Schemes of optical and electron microscopes</td>
      <td>152</td>
    </tr>
    <tr>
      <td>5.8</td>
      <td>Electron-matter interaction volume scheme.</td>
      <td>153</td>
    </tr>
    <tr>
      <td>5.9</td>
      <td>Batchs analysis: Collagen threads observed post-extrusion</td>
      <td>156</td>
    </tr>
    <tr>
      <td>5.10</td>
      <td>Viscosity as a function of shear rate for seven collagen batches at 5mg/mL.</td>
      <td>157</td>
    </tr>
    <tr>
      <td>5.11</td>
      <td>Collagen threads observed by different techniques at different concentrations.</td>
      <td>181</td>
    </tr>
    <tr>
      <td>5.12</td>
      <td>Evolution of collagen threads diameter over the time</td>
      <td>182</td>
    </tr>
    <tr>
      <td>5.13</td>
      <td>SEM images of threads surface at different concentrations</td>
      <td>183</td>
    </tr>
    <tr>
      <td>5.14</td>
      <td>Ultrastructure of threads after 2 weeks of maturation in PBS 5X as seen by TEM (transverse sections) at different concentrations</td>
      <td>184</td>
    </tr>
    <tr>
      <td>5.15</td>
      <td>P-SHG imaging of collagen threads at a.15 mg/mL, b. 30 mg/mL and c. 60mg/mL</td>
      <td>185</td>
    </tr>
  </tbody>
</table>
<table>
  <tbody>
    <tr>
      <td>5.16</td>
      <td>SHG imaging and P-SHG areas (colored) of surfaces and middle at the center thread transverse section</td>
      <td>186</td>
    </tr>
    <tr>
      <td>5.17</td>
      <td>SHG and P-SHG imaging of collagen thread at 30mg/mL (B) at day 1.</td>
      <td>187</td>
    </tr>
    <tr>
      <td>5.18</td>
      <td>SHG images of collagen threads at 60 mg/mL after 2 weeks in PBS 5X.</td>
      <td>187</td>
    </tr>
    <tr>
      <td>5.19</td>
      <td>Collagen:alginate threads at ratios: 1:1, 2:1, 5:1, observed by different techniques.</td>
      <td>189</td>
    </tr>
    <tr>
      <td>5.20</td>
      <td>SEM imaging of pure ALG4 (control) and collagen:alginate (5:1) threads.</td>
      <td>190</td>
    </tr>
    <tr>
      <td>5.21</td>
      <td>TEM longitudinal sections images of two characteristic areas of collagen:alginate (5:1) thread.</td>
      <td>191</td>
    </tr>
    <tr>
      <td>5.22</td>
      <td>TEM micrographs obtained from other studies to be compared with hybrid thread microstructure.</td>
      <td>191</td>
    </tr>
    <tr>
      <td>5.23</td>
      <td>SHG image of collagen:alginate (5:1) thread section and the respective projections.</td>
      <td>192</td>
    </tr>
    <tr>
      <td>5.24</td>
      <td>Tensile testing until rupture for collagen threads at 30 and 60mg/mL and coll:alg (5:1) thread in wet conditions and at room temperature.</td>
      <td>193</td>
    </tr>
    <tr>
      <td>5.26</td>
      <td>SEM images of collagen:alginate (5:1) thread surface after 3 weeks in DMEM at 37°C.</td>
      <td>195</td>
    </tr>
    <tr>
      <td>5.27</td>
      <td>Evolution of collagen based thread diameters upon maturation for the different buffers</td>
      <td>195</td>
    </tr>
    <tr>
      <td>5.28</td>
      <td>TEM cross-sections images of collagen threads at 30 mg/mL (B) and 60 mg/mL (C) either matured in PBS5X or incubated in DMEM at 37°C during 2 weeks.</td>
      <td>196</td>
    </tr>
    <tr>
      <td>5.29</td>
      <td>TEM sections along the thread axis (left //) and perpendicular (right ⊥) of collagen:alginate (5:1) threads incubated in DMEM at 37°C during 3 weeks.</td>
      <td>197</td>
    </tr>
    <tr>
      <td>5.30</td>
      <td>SHG image of collagen:alginate (5:1) thread section incubated 3 weeks in DMEM at 37°C.</td>
      <td>198</td>
    </tr>
    <tr>
      <td>5.31</td>
      <td>DSC measurements: denaturation of collagen 30mg/mL, coll:alg (5:1) and ALG4 threads kept either in their extrusion solutions (PBS 5X or CaCl<sub>2</sub> 1M respectively), or in DMEM during 2 weeks.</td>
      <td>199</td>
    </tr>
    <tr>
      <td>6.1</td>
      <td>Cell C3H/10T1/2 morphology.</td>
      <td>207</td>
    </tr>
    <tr>
      <td>6.2</td>
      <td>Protocol of cell culture under tension</td>
      <td>208</td>
    </tr>
    <tr>
      <td>6.3</td>
      <td>Scheme of collagen thread cell culture configuration under tension.</td>
      <td>208</td>
    </tr>
    <tr>
      <td>6.4</td>
      <td>Confocal Fluorescence Microscope scheme</td>
      <td>209</td>
    </tr>
    <tr>
      <td>6.5</td>
      <td>Fluorescence spectra of DAPI and Alexa Fluor 488</td>
      <td>210</td>
    </tr>
    <tr>
      <td>6.6</td>
      <td>In situ hybridization main steps</td>
      <td>211</td>
    </tr>
    <tr>
      <td>6.7</td>
      <td>Scheme of real time qPCR main steps</td>
      <td>212</td>
    </tr>
    <tr>
      <td>6.8</td>
      <td>Schemes of melting curves for the calibration points (top) and its derivative curve (bottom) with the melting temperature Tm.</td>
      <td>215</td>
    </tr>
    <tr>
      <td>6.9</td>
      <td>In situ hybridization main steps</td>
      <td>216</td>
    </tr>
    <tr>
      <td>6.10</td>
      <td>Tensile testing results of collagen threads at 30 mg/mL (blue) and 60 mg/mL (purple) seeded with cells and cultured over 3 weeks under static load and their respective controls.</td>
      <td>217</td>
    </tr>
    <tr>
      <td>6.11</td>
      <td>SHG images coupled with fluorescence of collagen threads seeded with C3H10 T1/2 cells during 3 weeks at 30mg/mL and 60mg/mL, under static load.</td>
      <td>219</td>
    </tr>
    <tr>
      <td>6.12</td>
      <td>SHG images coupled with fluorescence of cell sheath surfaces on collagen threads</td>
      <td>220</td>
    </tr>
    <tr>
      <td>6.13</td>
      <td>In situ hybridization on collagen threads at 30 and 60 mg/mL seeded with C3H10 T1/2 cells during 3 weeks</td>
      <td>221</td>
    </tr>
  </tbody>
</table>
<table><tr><td>6.14 Morphologies observed by phase contrast microscopy of collagen thread at 30 mg/mL (left) and collagen:alginate (5:1) thread (right) after being seeded with cells and cultured during 3 weeks</td><td>223</td></tr><tr><td>6.15 Confocal microscopy images of collagen thread at 30 mg/mL (left) and mixture coll:alg (5:1) (right) for different times: days 3, 7, 21 and without tension at 21 days.</td><td>225</td></tr><tr><td>6.16 SHG image coupled with fluorescence of coll:alg (5:1) thread at 2 weeks of culture with cells without static load.</td><td>226</td></tr><tr><td>6.17 In situ hybridization on collagen threads at 30 mg/mL (left) and coll:alg (5:1) thread (right) with C3H10 T1/2 cells at 3 weeks, under static load.</td><td>227</td></tr></table>
# List of Tables

<table><tr><td>1.1</td><td>Mechanical properties of human tendons</td><td>12</td></tr><tr><td>2.1</td><td>Characteristic dimensions of stainless steel needles</td><td>49</td></tr><tr><td>2.2</td><td>Persistence length for different buffers found in the literature.</td><td>75</td></tr><tr><td>3.1</td><td>Power law fit exponent values of viscosities for collagen at 30 mg/mL</td><td>91</td></tr><tr><td>3.2</td><td>Characteristic physical quantities of collagen solutions of interest</td><td>94</td></tr><tr><td>3.3</td><td>Characteristic physical quantities of chosen alginates solutions.</td><td>96</td></tr><tr><td>3.4</td><td>Fit exponents α and β for the respective storage and loss moduli; the crossover angular frequency ω<sub>c</sub> and the relaxation time λ</td><td>97</td></tr><tr><td>5.1</td><td>Characteristic physical quantities of collagen solutions of interest</td><td>193</td></tr><tr><td>5.2</td><td>Characteristic physical quantities of collagen solutions of interest after incubation in DMEM</td><td>200</td></tr><tr><td>6.1</td><td>Table of the primers</td><td>214</td></tr></table>