Skin Bioprinter Design and Construction

Tissue engineering has had initial successes with building a number of tissues clinically, however, challenges still exist in developing and translating complex tissue systems that require coordinated and systemic function. Based on the shortcomings of the current technologies and therapies of wound treatments, we established design criteria for an in situ skin bioprinting system. The bioprinting system is: (1) portable and capable of being quickly transported, (2) able to identify and measure a wide range of wound sizes and topologies accurately, (3) capable of delivering multiple cell types to precise spatial orientation tailored to a individual wound, (4) easily sterilized, and (5) is easily operated and maintained with relatively low cost. A schematic demonstrating scale, design and components of the bioprinter system is shown in Fig. 1A. The main components of the system consist of a hand-held 3D wound scanner, and a print-head with an XYZ movement system containing eight 260 µm diameter nozzles, each driven by an independent dispensing motor. All components are mounted on a frame small enough to be mobile in the operating room (Fig. 1B). The dimensions of the system are 79 cm wide (patient head to toe direction) by 77 cm deep (cross patient direction). When the robotic arm is fully extended, it hands out over the base an additional 50 cm so the full reach is 127 cm, which is large enough to cover the torso of an average patient but small enough to easily pass through most doorframes. The skin bioprinter system is equipped with locks that engage with the base of the patient table to prevent movement while printing cells. Once the system is in position over the patient these locks restrain the mobility of the system during the printing process to ensure high delivery precision. The XYZ axes contribute to delivery precision by using a plotting system capable of 100 µm movements. In addition, the Z-axis allows the delivery system to maintain the set distance from the surface of the wound to accommodate the curvature of patient’s body. Reconstructing skin wounds can be tailored to an individual patient by combining a wound scanning system with the cartridge-based delivery system (Fig. 1C). To print a skin construct that exactly matches a patient’s wound, the current prototype incorporates a real-time 3D laser scanner, ZScanner™ Z700 scanner (3DSystems, Rock Hill, SC), utilizing markers that are placed around the wound area used as reference points (Fig. 1D). The hand-held scanner is easy to use and allows for easy maneuverability. The scanner offers the ability to capture the entire wound in one continuous scan. Once the scan is complete, the wound data is compiled to form a model of the wound. The scanned wound area is then processed using Geomagic Studio software (Morrisville, NC) orient the model to standard coordinates so the import into Artcam is ideal. Scanned data is in the form of an STL file that has been outputted from Geomagic and inputted to Artcam® 3D software to obtain the full volume and the nozzle path needed to print the fill volume. The wound depth is then split into layers on the Z axis to determine which layers correspond to the dermis and epidermis (Fig. 1E). Each layer is overlaid with a series of XY lines that cover the entire wound area. These lines are used in conjunction with the cartridge-based delivery system to determine a path for the printer. The skin cell delivery system is controlled by a custom software employing a three-tiered architecture design based on the Microsoft.NET Framework 2.0 (Microsoft Corp®. Redmond, WA) and written in C++. The three-tiered architecture handles three areas of communication: between the user and the software, among the software components, and between the software and the file system. This design structure allows every component to be quickly and easily altered as necessary without affecting the other components.

Figure 1 Skin bioprinter prototype and in situ bioprinting concept. (A) Schematic demonstrating scale, design and components of the skin bioprinter. (B) The main components of the system consist of 260 µm diameter nozzles, driven by up to 8 independently dispensing systems connected to a print-head with an XYZ movement system, in addition to the 3D wound scanner. All components are mounted on a frame small enough to be mobile in the operating room. (C) Skin bioprinting concept. Wounds are first scanned to obtain precise information on wound topography, which then guides the print-heads to deposit specified materials and cell types in appropriate locations (Images courtesy of LabTV - National Defense Education Program, Washington, D.C.). (D) Example of skin bioprinting process, where markers that are placed around the wound area used as reference points (a) prior to scanning with a hand-held ZScanner™ Z700 scanner (b). Geometric information obtained via scanning is then inputted in the form of an STL file to orient the scanned images to standard coordinate system (c). The scanned data with its coordinate system is used to generate the fill volume and the path points for nozzle head to travel to print the fill volume (d). Output code is then provided to the custom bioprinter control interface for generation of nozzle path needed to print fill volume (e,f). (E) This system facilitates the depositing of multiple cell types with high precision and control. Layering of fibroblasts (green) and keratinocytes (red) is shown. Full size image

The cartridge-based delivery system is similar to that used in traditional inkjet printing. The pressure-based delivery system uses three components: a delivery mechanism, the material reservoirs and a pressure source. The delivery system is based on cartridges that can contain any material that can be delivered through the print-head nozzles, with each nozzle connecting to a separate cartridge. For the skin delivery system the contents of each cartridge are embedded in a matrix composed of fibrinogen and collagen. Separate atomizing nozzles simultaneously deposit thrombin on the fibrinogen matrix to produce fibrin. Each cell type is loaded into an individual cartridge in the same way different color inks would be contained in different cartridges. For our clinical printer, design updates were performed to incorporate a more controllable piston-driven pressure mechanism. For the air-pressure system, air from the pressure source fills the reservoir and drives the material in the reservoir into the delivery mechanism as it is needed. The system is pressurized with 6.89 kPa of compressed air using a two-stage control valve. The piston-driven pressure delivery was achieved with individual syringe pumps, and was able to achieve identical pressures but with more control over ramping and depressurization. Both systems resulted in identical functional outcomes. Together, these components provide a powerful new tool for treatment of large wounds.

Bioprinting Procedure

The sterilization, preparation and bioprinting procedures are easy and quick to perform. The system is designed to be covered with a sterile drape. The print-head is detachable and able to be autoclaved. Sterilization of tubing was performed by connecting cartridges of 70% ethanol followed by nano-filtered autoclaved water to clear the residual ethanol. A sterilization command directs the printer to automatically flush the delivery system with ethanol for 3 minutes followed by a sterile water flush for 1 minute. The user can choose to print using one of three different methods. First, the laser scanner can be used as previously described to create a wound map for printing. Second, the user can define a series of bitmap images where each non-black pixel corresponds to a cell drop, and the color of the pixel corresponds to the cell type to print. This method allows the user to create complex structures using any cell type in any configuration. Finally, a list of available cell types loaded in the printer is provided to the user by the print-head configuration. In each method, the user defines the spacing between drops, and this is only limited by the resolution of the motors in the plotting system. A laser sight guided the user to the upper-left corner of the wound and the distance from the laser to the first inkjet valve was used as a calibration that ensured accurate and repeatable delivery of cells. The fibrin/collagen hydrogel carrier used in this study provides suitable support to maintain cellular viability, rapid crosslinking, and accurate depositing to facilitate the formation of multicellular, multi-layered skin constructs. In these studies the integrated scanning system enabled us to rapidly measure and convert wound size, depth and topology into a two layered format comprising of a lower fibroblast layer with an upper keratinocyte layer.

Bioprinter Validation in a Murine Full-Thickness Wound Model

We first validated the in situ skin bioprinting system using a murine full thickness excisional wound model. In the 36 female outbred athymic nude (Nu/nu) mice used in this study, the mortality rate was 0% and no wound infection or major skin irritation was noted on any mouse at any time point. Gross examination revealed that every printed skin construct appeared to form an epithelial coverage over the wound at 1 week (Fig. 2A). This epithelium became more organized with time until the wound was completely covered. Organization into fully-formed skin required approximately 10–14 days post-printing.

Figure 2 Gross examination of printed skin in a murine full thickness excisional wound model. (A) Cell printed mice show epithelium forming over the wound as early as week 1, with developing skin observed at week 2 that covers the entire wound but has not fully formed. By week 3, cell printed mice show complete coverage of the wound. Between week 4 and week 6, minimal contraction is observed and we observe a maturing epithelium. In contrast, matrix-treated and untreated wounds show minimal epithelialization until week 4, resulting in a significant proportion of open wound area. Contraction is also significant between weeks 4 and 6 following closure of the wounds. Scale bar: 1 cm. (B) Analysis of murine wound sizes over 6 weeks. Printed skin constructs show a significantly reduced time to wound closure when compared to untreated and matrix-treated controls. Printed skin closed the wound in 3 weeks compared to 5 weeks for controls. Wound sizes were analyzed with one-way ANOVA. ****p < 0.0001, n = 12; ***p < 0.01, n = 8; *p < 0.05, n = 8. Data presented as mean ± standard deviation (SD). (C) Anti-human nuclear antigen showed printed skin at the center of the wounds contained human within the epidermis and dermis at week 3, and week 6 after bioprinting. Matrix printed control wounds showed no human cells present at 6 weeks. Scale bar: 100 µm. (D) Masson’s Trichrome examination of skin. (a–c) Printed skin at the center of the wound shows increased cellularity compared to the matrix-treated and untreated controls, but lacked the presence of a defined epidermis or dermis maturation. At week 3 and week 6 post-printing we observed the presence of a defined epidermis and organized dermis consisting of aligned blue stained collagen fibers. (d–f) Matrix-printed wounds lacked cellularity and presence of a defined epidermis or dermis maturation one week after printing. At week 3 after treatment, matrix-printed still lacked a defined epidermis and showed no dermal organization but started to show epidermis formation and a more defined dermis with increased prominence of blue stained collagen fibers by week 6 (g–i) Untreated wounds lacked both the cellularity and material thickness observed in the other two groups at week one, but appeared similar to matrix only-treated wounds at week 3 and 6. Scale bar: 100 μm. Full size image

Wounds treated using in-situ skin bioprinting demonstrated faster wound closure compared to untreated and matrix-treated group (Fig. 2B). In the printed group, the wound area at 1 week post-surgery was 66% of the original wound area. In contrast, the wounds in the control groups remained at 95% of their original wound size at the same time point (n = 12, p ≤ 0.001). At 2 weeks, the wounds treated with printed cells were only 15% of the original size, while the wounds in the control groups were 40% of the original size (n = 8, p ≤ 0.05). Overall, printed skin cells were able to close the entire wound by 3 weeks post-surgery compared to 5 weeks for both negative controls.

Immunohistochemistry for human cells showed that human fibroblasts and keratinocytes were present within the dermis and epidermis of the wound 3 weeks and 6 weeks after printing, together with endogenous cells (Fig. 2C). Control groups did not show any staining for human cells. Histological examination of retrieved skin constructs revealed high cellularity of printed wounds one week after printing (Fig. 2D). This was expected, as our system delivers approximately 500 cells per drop using the current prototype. This histological finding supports the observation of epithelialization of the bioprinted wounds, and indicates that cells were present in the wound, perhaps forming an immature epidermal barrier. At week 3 and week 6 post-printing we observed the presence of a defined epidermis and organized dermis consisting of aligned blue stained collagen fibers. This finding is consistent with the accelerated wound closure and re-epithelialization. In matrix-printed wounds, the deposited hydrogel was visible within the wound area one week after printing, but as expected, lacked cellularity. At week 3 after treatment, matrix only-treated wounds showed increased cellularity, suggesting migration of endogenous cells, however these wounds lacked a defined epidermis and showed minimal dermal organization. At week 6 after treatment matrix-only treated wounds started to show epidermis formation and a more defined dermis with increased prominence of blue stained collagen fibers. Untreated wounds lacked both the cellularity and material thickness observed in the other two groups at week one, but appeared similar to matrix only-treated wounds at week 3 and 6 In all printed constructs the dermis contained a highly organized deep reticular layer and a less-organized papillary layer. The major three layers of the skin were present. All groups showed complete wound closure and healing by the end of week 6.

Bioprinter Validation in a Porcine Full-Thickness Wound Model

An overview of the porcine model study design is shown in Supplementary Fig. S1. All wounds formed granulation tissue by week 1 following excision, with greater levels of granulation appearing in the matrix, allogeneic and autologous cell-treated wounds (Fig. 3A). By week 2 we observed granulation tissue still apparent in the allogeneic and autologous cell-treated wounds, but to a lesser extent in the matrix-treated wounds. At this time, we also observed the beginning of contraction in the untreated wounds, with tattoo borders beginning to reduce in size and curve inwards, presenting a typical contractile wound shape. Autologous cell-treated wounds appeared to form a thin epithelium over the majority of the wound area by 2 weeks, which then thickened and became white opaque from 4 weeks onwards. Interestingly, these opaque, white areas of epithelium originated from small patches within the wound area, and grew throughout the time-course of the experiment, suggesting that these epithelial patches did not originate from the tissue edge but rather from autologous cells applied with the bioprinter. During weeks 2 to 4, autologous cell-treated wounds contracted very little, while all other treatments appeared to contract significantly, becoming reduced in total size, but also losing the square shape and becoming rectangular with pointed corners, suggesting contraction toward the center of the wounds. These trends continued between weeks 4–8, with autologous cell-treated wounds showing increased opaque, white-appearing epithelium, while maintaining tissue shape and size. In contrast other groups continued to contract, with little epithelium forming primarily from the edges of the wound. Between weeks 6 and 8 it appeared that all wounds had reached their final contracted size.

Figure 3 Gross examination of printed skin in porcine model. (A) Images of wound healing of in situ bioprinted autologous and allogeneic fibroblasts and keratinocytes compared to bioprinted fibrinogen/collagen (matrix only) and untreated control groups over 8 weeks. Formation of epidermis islands at the core of the autologous treatments started as early as week 2 and complete wound closure and re-epithelialization with minimal contracture was achieved by week 4 of the study. All other treatments appeared to contract significantly, with little epithelium forming primarily from the edges of the wound. (B) Significantly smaller open wound size was measured for autologous cell-treated wounds compared to other treatments. (C) Wound contraction was also reduced in autologous cell-treated wounds compared to other treatments. (D) Autologous cell-treated wounds showed significantly accelerated re-epithelialization compared all other groups. Wound sizes were analyzed with one-way ANOVA. ***p < 0.001, n = 6; **p < 0.01, n = 6; *p < 0.05, n = 6. Data presented as mean ± standard deviation (SD). Full size image

Wound size was measured for each wound once per week and recorded as area in centimeters squared (cm2) (Fig. 3B). One week following surgery, most wounds maintained their original size, however, contraction did occur in the untreated wounds, resulting in a decrease in wound size to a mean of 87.75 ± 15.25 cm2 (p < 0.05). From this time-point onwards, untreated wounds were statistically indistinguishable in size from matrix, and allogeneic cell-treated wounds, although the trend was for a smaller wound size at weeks 1 to 3. From week 2, 3, 4 and 5 autologous cell-treated wounds closed rapidly, and open wound area as significantly smaller than all other groups. Specifically measuring: 16.14 ± 15.29 cm2 at week 2 (p < 0.001 vs all groups), 3.40 ± 4.64 cm2 at week 3 (p < 0.001 vs all groups), 1.46 ± 2.09 cm2 at week 4 (p < 0.001 vs all groups), and 0.00 ± 0.00 cm2 at week 5 (p < 0.05 vs all groups). From this point onwards other treatments also trended in this direction, with no statistical differences between groups. These data suggest that the in situ bioprinting of autologous cells resulted in approximately 3-week acceleration in wound closure compared to other treatments.

Wound contraction was measured for each wound at the wound tattoo border once per week and expressed as a percentage of the original wound tattoo area (Fig. 3C). One week after treatment, all wounds other than untreated expanded in size, resulting in a negative contraction value. Untreated wounds only showed minimal expansion. This effect is observed in many of our excisional wound models, and represents a combination of sagging or stretching of the wound area following full thickness excision, with some contribution from swelling and granulation. By week 2, this trend was reversed with all groups showing positive contraction (shrinking), without statistically significant differences between groups. At week 3 autologous cell-treated wounds were significantly less contracted than untreated wounds (14.33 ± 5.99% vs 34.3233 ± 10.24%, p < 0.05), and at week 4 autologous cell-treated wounds were significantly less contracted than matrix-treated wounds (20.00 ± 8.96% vs 39.91 ± 12.42%, p < 0.05). However compared to allogeneic-treated wounds there were no statistical differences at these time-points, although the obvious trend for the duration of the study was reduced contraction in autologous cell-treated wounds. From weeks 5 to 8, autologous cell-treated wounds showed significantly less contraction than both untreated and matrix-treated wounds (week 5, p < 0.05; week 6–8, p < 0.01) and also less contraction than allogeneic cell-treated wounds (p < 0.05). These data suggest that bioprinting autologous cells to full thickness wounds resulted in approximately 50% reduction in wound contraction compared to other treatments over the time-course of the study.

Wound re-epithelialization was expressed as a percentage of total area within the tattoo border covered by epithelium (Fig. 3D). No epithelialization was recorded at week 0 or week 1 for any group. As early as week 2, autologous cell-treated wounds showed significantly greater percentage of re-epithelialization compared to all groups between weeks 2 and 5. Specifically measuring: 86.83 ± 13.30% at week 2 (p < 0.001 vs all groups), 93.73 ± 7.12% at week 3 (p < 0.001 vs all groups), 97.24 ± 4.23% at week 4 (p < 0.001 vs all groups), and 97.00 ± 6.71% at week 5 (p < 0.001 vs all groups). At week 6, autologous cell-treated wounds showed significantly greater percentage of re-epithelialization than both untreated and matrix-treated wounds (98.57 ± 3.50%, p < 0.05), but this was not significantly different to allogeneic cell-treated wounds (92.11 ± 6.17%). Similar to the wound closure data, from this point onwards, autologous cell-treated wounds showed complete wound closure and re-epithelialization and other treatments trended in this direction, with no statistical differences between groups. These data show that bioprinting autologous cells to full thickness wounds resulted in rapid epithelialization of the wounds, representing a 4–5 week acceleration in wound re-epithelialization compared to other treatments.

Porcine Wound Histological Analyses

Microscopic examination identified increasing degrees of tissue regeneration, epithelialization and maturation as wound healing progressed through the weeks (Fig. 4). All wounds progressed through an initial formation of granulation tissue on the surface followed by a thin discontinuous layer of squamous epithelium that eventually became continuous and covered the entire wound. Wounds receiving bioprinted autologous cells showed early epithelialization with almost complete coverage of the wound at 2 weeks. By week 4, autologous bioprinted wounds showed the formation of rete peg epithelial projections into the dermis, as well as the presence of keratinized stratified squamous epithelium. In contrast, allogeneic cell and matrix-treated wounds did not show the presence of epithelialization or keratinized stratified squamous epithelium until week 6. Untreated wound healing appeared even more delayed, with only an immature epithelium present at week 6, which still lacked the mature-appearing rete peg epithelial projections and keratinized stratified squamous epithelium at week 8.

Figure 4 Microscopic examination of H&E stained sections showed increasing degrees of tissue regeneration, epithelialization and maturation as wound healing progressed. Wounds receiving bioprinted autologous cells showed early epithelialization with almost complete coverage of the wound at 2 weeks. By week 4, autologous bioprinted wounds showed the formation of rete peg epithelial projections into the dermis, as well as the presence of keratinized stratified squamous epithelium. Wounds treated with bioprinted autologous cells showed early formation of a loosely organized papillary layer above a denser, thicker layer of reticular dermis. The structure of the tissue appeared mature and complete at 8 weeks with nicely woven dermal collagen and regularly distributed vasculature. Magnification 20x, Scale bars 100 µm. Full size image

Wounds treated with bioprinted autologous cells showed early formation of a loosely organized papillary layer above a denser, thicker layer of reticular dermis. The papillary dermis appeared to consist of coarse bundles of larger loose fibers arranged parallel to the epidermis, while the reticular dermis contained larger blood vessels and closely interlaced smaller fibers. The structure of the tissue appeared mature and complete at 8 weeks with nicely woven dermal collagen and regularly distributed vasculature. Allogeneic cell and matrix-treated wounds showed the continued presence of granulation tissue 2 weeks after treatment, progressing to a dermis consisting of a densely packed fibroblastic proliferation lacking distinction between the papillary and reticular dermal layers at week 4. By weeks 6 and 8 the dermis in these groups appeared similar to the autologous cell-treated group in appearance, matching the wound healing progression time frame observed for the epidermis. Untreated wounds generally lacked the granulation tissue formation observed histologically for other groups, showing only a densely packed dermis and lacking blood vessel formation. This continued until week 6, where the dermis matured at a similar, but slower progression than seen with other groups.

It is well known that the more prolonged the inflammatory phase of wound healing is, there is more likelihood of generating a non-functional scar. Therefore, we focused on assessing the degree and types of inflammation histologically. A pathologist blinded to the treatment groups scored acute and chronic inflammation in H&E stained sections using the grading criteria shown in Supplementary Fig. S2. Acute inflammation was most abundant in week 2 samples of all groups and decreased over the wound healing progression (Table 1). Acute inflammation was mostly superficial and appeared to be inversely related to the degree of epithelialization. It was very prominent in all groups at 2 weeks (score of 3) except for the autologous cell-treated wounds, which only showed mild acute inflammation (score of 1). Acute inflammation was mostly absent or very mild by the 8th week in all groups. Chronic inflammation was present in all wounds and it was mainly in the deeper aspects of the wound. It persisted throughout the study duration and did not appear to affect maturation and re-epithelialization of the wound. Granulomatous inflammation and foreign body giant cells were noted in the deeper aspects in some of the grafts with no noticeable association with any specific treatment type.

Table 1 Grading of acute and chronic inflammatory response of treatments. Full size table

To evaluate the formation and maturation of wound vasculature, CD31 immuno-staining was performed at weeks 4 and 8 of wound regeneration (Fig. 5A). Concurring with our observations of early granulation tissue formation in the autologous cell-treated wounds, we identified a higher density of CD31-positive blood vessels throughout the dermis of these wounds, with only minimal, small blood vessels sparsely distributed throughout the dermis of the other groups. By week 8, the overall number of small blood vessels appeared to decrease in autologous cell-treated wounds, and instead we observed larger mature vessels throughout the dermis. In contrast, untreated wounds showed large numbers of small, immature blood vessels, suggesting that these tissues were undergoing delayed wound healing, appearing similar to autologous cell-treated wounds at week 4. Matrix and allogeneic cell-treated wounds lacked the high density of small blood vessels observed in the week 4 autologous, and week 8 untreated wounds, and instead showed a reduced number of mature blood vessels.

Figure 5 Histological evaluation of vascularization, collagen deposition, myofibroblast activation, and cell proliferation. (A) We observed a higher density of CD31-positive blood vessels throughout the dermis of autologous cell-treated wounds at week 4, transitioning to larger mature vessels throughout the dermis at week 8. (B) Blue stained collagen fibers were most prominent in matrix-treated and autologous cell-treated wounds. Collagen fibers present in the autologous cell-treated wounds appeared much larger and organized than observed in other groups. (C) αSMA-positive cells were more prominent in the autologous cell-treated wounds at week 4. At week 8, the untreated and matrix-treated wounds showed significantly greater numbers of αSMA-positive cells, distributed throughout the dermis, and surrounding the sparse blood vessels, suggesting the presence of large numbers of contractile myofibroblasts at this time-point. (D) At week 4, untreated, matrix and allogeneic cell-treated wounds showed the greatest number of proliferating cells, primarily located throughout the dermis. Autologous cell-treated wounds showed fewer overall proliferating cells, however these cells were mostly present immediately underlying the developing epidermis, suggesting that these cells were keratinocytes contributing to epidermis formation and maturation. CD31/Ki67: Magnification 20x, Scale bars 100 µm, Trichrome/αSMA: Magnification 40x, Scale bars 200 µm. Full size image

Masson’s trichrome staining of tissue sections at week 4 highlighted a dermis composition with densely packed nuclei, potentially due to inflammatory cells, and generally lacking organized collagen fibers (Fig. 5B). This was most prominent in the untreated and matrix treated groups. Allogeneic and autologous cell-treated groups showed more diffuse nuclei distributed in a background of organized collagen fibers. At week 8, the concentration of cell nuclei was significantly reduced in all groups, however patches of concentrated cells were still present in the dermis of the untreated, matrix-treated and allogeneic cell-treated groups. These were still present but fewer in number in autologous cell-treated groups. Blue stained collagen fibers were most prominent in matrix-treated and autologous cell-treated wounds. Interestingly this was not observed to the same extent in allogeneic cell-treated wounds. Collagen fibers present in the autologous cell-treated wounds appeared much larger and organized than observed in other groups, supporting histological observations that the dermis of these wounds appeared more mature than for other treatments.

Fibroblastic differentiation into myofibroblasts plays a major role in the formation of wound granulation tissue early in wound healing, and persists in contractile wounds, such as hypertrophic scars. To evaluate the presence of myofibroblasts in the healing wounds, we performed immuno-histochemistry for α-smooth muscle actin (αSMA) at weeks 4 and 8 (Fig. 5C). αSMA-positive cells were largely absent from untreated and matrix-treated wounds at week 4, while the allogeneic and autologous cell-treated wounds showed moderate staining in the papillary region of the dermis. This staining was more prominent in the autologous cell-treated wounds. At week 8, the untreated and matrix-treated wounds showed significantly greater numbers of αSMA-positive cells, distributed throughout the dermis, and surrounding the sparse blood vessels, suggesting the presence of large numbers of contractile myofibroblasts at this time-point. In the allogeneic and autologous cell-treated wounds, αSMA staining was primarily located around the larger mature blood vessels, supporting observations of increased blood vessel formation and maturation in these tissues.

Ki67 immuno-staining was performed at weeks 4 and 8 to evaluate the presence of proliferating cells within the wounds (Fig. 5D). At week 4, untreated, matrix and allogeneic cell-treated wounds showed the greatest number of proliferating cells, primarily located throughout the dermis, potentially activated myofibroblasts, and at the wound surface, which had not yet formed an immature epidermis. Autologous cell-treated wounds showed fewer overall proliferating cells at week 4, however these cells were mostly present immediately underlying the developing epidermis, suggesting that rather than proliferating myofibroblasts, these cells were keratinocytes contributing to epidermis formation and maturation. At week 8, the untreated, matrix and allogeneic cell-treated wounds showed a similar staining pattern to that observed in the week 4 autologous cell-treated wounds, while the week 8 autologous cell-treated wounds showed an even greater number of proliferating cells in the underlying epidermis area.

Skin bioprinting comparison with cell spraying

We evaluated the efficacy of our bioprinting technique and compared to a cell spraying technique, which is becoming commonly applied in the clinic for treatment of wounds using autologous skin cells. Specifically, we compared our in situ skin bioprinting system using the printing parameters described above, to the FibriJet biomaterial delivery device using the same materials, concentrations, volumes and application timing for both techniques. Similar to our previous studies, both groups formed granulation tissue by 2 weeks followed by rapid wound closure through a combination of contraction and re-epithelialization, resulting in total wound closure (Supplementary Fig. S3A). Although the bioprinted wounds appeared to show more rapid wound closure, reduced contraction and increased re-epithelialization, there were no statistical differences for these parameters at any time-points (data not shown). This suggests that for these parameters, our bioprinting system is equally as effective as the currently available clinical cell spraying technologies.

Interestingly, histological analysis of the wounds at each time-point showed that the bioprinted wounds resulted in earlier epithelialization with almost complete wound coverage at 2 weeks, while the cell sprayed wounds did not show the formation of epidermis until week 4 (Supplementary Fig. S3B). In fact, the epidermis of the cell spray groups did not thicken and showed signs of maturation such as presence of keratinized stratified squamous epithelium until 6 weeks. Wounds treated with the bioprinted cells showed early formation of a papillary layer consisting of coarse bundles of larger loose fibers, distinct from a dense, thicker layer of reticular dermis containing larger blood vessels and closely interlaced smaller fibers. In contrast, wounds treated with the sprayed cells showed a dermis consisting of a densely packed organization lacking a distinction between the papillary and reticular dermal layers at week 2, which eventually matured to form a more distinct and mature dermis similar to bioprinted wounds by week 4. Trichrome staining supported these observations, confirming that the bioprinted wounds showed accelerated formation of epidermis and more mature dermis tissue at weeks (Supplementary Fig. S3C). Collagen fibers stained in blue were more prominent in the bioprinted wounds at week 4, compared to wounds treated with the sprayed cells. By week 6, both groups had similar appearance with a defined epidermis and organized dermis, consisting of aligned collagen fibers.