The classical approach for treating skin loss due to disease or injury is autologous grafting (Table 1), where skin is removed from the patient at an alternative site to assist with closure of the primary wound (Herskovitz et al., 2016). Grafts can be classified as full-thickness or split-thickness, where full-thickness includes the entirety of the epidermis and dermis, and split-thickness include epidermis and only part of the dermis (Herskovitz et al., 2016). The clinical gold standard remains an autologous, split-thickness skin graft. Despite their popularity, skin grafts can be problematic because they involve production of another wound at a secondary site, placing further healing burden on the patient (Dearman et al., 2021). Skin grafts come with additional risks, including rejection, infection, and seroma (Kohlhauser et al., 2021). The donor site is particularly problematic, with moderate to high reported incidences of pain, hypertrophic scarring incidence up to 28%, infection, and reduced quality of life (Asuku et al., 2021).

In cases where sufficient tissue cannot be provided by the patient, allogenic grafts, where the graft is harvested from an alternative donor, can be used (Table 1) (Schlottmann et al., 2021). Allogenic grafts come with a greater risk of immunogenicity and subsequent graft rejection due to their foreign nature (Schlottmann et al., 2021). The healing outcomes following skin grafts are not always consistent, particularly following allogenic grafting (Schlottmann et al., 2021).

In the absence of autologous or allogenic skin grafting, skin substitutes can be used (Table 1) (Halim et al., 2010). These do not necessarily have to be human-derived but must be biocompatible and capable of supporting the wound as it heals. Using biomaterials as a base to construct porous or fibrous scaffolds is one strategy, producing 3-dimensional (3D) skin substitutes which provide structural support to the wound (Sheikholeslam et al., 2018). Biomaterials can be isolated from natural and synthetic sources (Lee et al., 2018). Biomaterial-based scaffolds can be produced by a number of techniques including self-assembly (Webber and Pashuck, 2021), electrospinning (Rahmati et al., 2021), 3D printing (Richards et al., 2013), and decellularisation (Neishabouri et al., 2022). Porosity of the scaffold is critical, because this can alter cell interactions and change the capacity for the scaffold to transfer nutrients and oxygen (Loh and Choong, 2013). The ECM that surrounds and supports skin cells in vivo is a complex arrangement of collagen, elastin, and glycosaminoglycans (GAGs), and each of these play a role in maintaining skin homeostasis and structural support, as well as contributing to the healing process following skin injury (Rousselle et al., 2019). Therefore, constructing skin substitutes from human ECM components, e.g., collagen, is a logical approach. Collagen molecules are triple helices that assemble into fibrils, a few hundred nanometres in diameter, which form basket weave structures within native skin (Fratzl, 2003). Mammalian collagen (e.g., from bovine, porcine, ovine sources) can be formulated into porous scaffolds by 3D-printing, and fibrous scaffolds by self-assembly or electrospinning (Sheikholeslam et al., 2018). As a biomaterial, collagen can interact with cells and the ECM of the recipient to promote cellular proliferation (Parenteau-Bareil et al., 2010). Mammalian collagen scaffolds include Bio-ConneKt and HeliColl, among others (Table 1) (Sharma et al., 2023). Unfortunately, mammalian collagen has been associated with immunogenicity following implantation, with ethical controversy complicating its use (Parenteau-Bareil et al., 2010).

Decellularisation allows for preservation of the ECM structure as a whole, isolating all of its components (collagen, elastin, GAGs) while removing potentially immunogenic cellular components (Crapo et al., 2011). Human and animal-derived decellularised matrices are commercially available for wound healing indications (Table 1), including Alloderm^® and GraftJacket™ from donor cadaver skin (Dussoyer et al., 2020) and Matristem urinary bladder matrix (UBM)™ from porcine bladder tissue (Kimmel et al., 2010). Placement of these structures on the wound allow for migrating cells from the neighbouring tissue to populate the scaffold as healing occurs (Wei et al., 2002). However, these structures are associated with complications. In addition to continued ethical criticism, mammalian acellular matrices can be inconsistent in structure and recellularisation capacity, based on the features of the donor (Gilpin et al., 2014; Johnson et al., 2016). Additionally, mammalian ECM components or ECM fragments generated during the decellularisation process are capable of inducing immune reactions within the recipient (Allaire et al., 1997; McQuitty et al., 2020). Production of these fragments can be linked to the chemical-heavy approaches currently used for decellularisation (Crapo et al., 2011).

Biologically inert materials are potentially advantageous over biologically active materials because they reduce the opportunity for immunogenicity while still providing the structural support required for healing (Hickey and Pelling, 2019). For example, cellulose is an inert biomaterial derived from bacterial and plant sources and is associated with good biocompatibility with wound healing cells (Modulevsky et al., 2014; Modulevsky et al., 2016; Koivuniemi et al., 2020). While cellulose molecules differ from those of collagen, in that they consist of chains with repeats of two sugar rings, they also naturally exist as thin, spiraling microfibrils, but contain nano-to micro-metre crystals depending on their source (Fratzl, 2003). Hydroxyl groups present on cellulose chains allow for cell adhesion and cellulose has high stability and tensile strength (Naomi et al., 2020). Cellulose can be extracted as crystals, fibres, or decellularised scaffolds, formulated as hydrogels, or used as bioink for 3D-printing (Sharip and Ariffin, 2019). Carboxymethylcellulose, a form of modified cellulose, is a well-characterised biomaterial in the wound space and has been formulated into products such as the AquaCel^® Hydrofiber dressing (Table 1) (Williams, 1999; Guest and Ruiz, 2005). Other biologically inert products for wound repair include the synthetic polyurethane scaffold Novosorb^® biodegradable temporising matrix (BTM), which has been highly successful in the treatment of complex wounds (Table 1) (Li et al., 2021). The production of these materials generally requires chemically-intensive processes, which can reduce their environmental sustainability (Radotić and Mićić, 2016; Gowthaman et al., 2021; Khan et al., 2022).

Incorporating cells into skin substitutes allows for synthesis of living grafts, which not only provide structural support to wounds, but contain functional skin cells that can contribute to the healing process (Table 1). Such cells include keratinocytes, fibroblasts, and endothelial cells. Keratinocytes are found throughout the epidermal layer of the skin where they create four to five differentiated layers from the deepest stratum basale to the outer stratum corneum (Montagna, 2012). The highly replicative keratinocytes within the stratum basale are attached to a collagen-based basement membrane below which lies the dermis. As keratinocytes transition through the epidermal layers, they increase in keratin and lipid production, then flatten and die to create the dry, protective barrier of the stratum corneum. Also found within the epidermis are melanocytes, Merkel cells and Langerhans cells, which respond to ultraviolet radiation, physical and microbial cues, respectively. Below, the dermis is heavily populated with fibroblasts, responsible for secreting collagen, elastin and GAGs which form the surrounding ECM and provide structure, tensile strength and elasticity, and endothelial cells, which comprise the vasculature and control oxygen and nutrient supply (Montagna, 2012). The dermal layer also contains phagocytes and lymphatic vessels critical for the response to microbial breaches of the skin. The innermost hypodermis, or subcutaneous layer, comprised of well vascularised, adipose tissue within loose ECM, which provides insulation and cushioning. The skin also comprises a number of accessory organs, including sebaceous glands, hair follicles, and nervous innervation (Montagna, 2012; Abdo et al., 2020). The maintenance and regeneration of cells and organs within the skin is directed by growth factors and signalling molecules, which must be supplied exogenously when constructing skin substitutes (Sharma et al., 2019).

The most simplified version of living grafts are cell sheets that secrete their own ECM. These cells are generally isolated from skin biopsies, from the patient or healthy donors. The isolated cells are expanded in vitro through the addition of growth supplements, then applied as thin films of confluent cell layers (Chaudhari et al., 2016). Further, autologous self-assembled skin substitutes (SASS) have been created that allow for replacement of both the dermis and epidermis in a single procedure. An acellular dermis construct is prepared through self-synthesis of a collagen-rich ECM by the patient’s fibroblasts. The patient’s keratinocytes are then cultured on the dermal construct to form a stratified epidermis, which is transplanted onto full-thickness burns or wounds (Athanasiou et al., 2013; Dagher et al., 2023). In addition, spray suspensions of keratinocytes and fibroblasts have been used successfully to apply cells in a thin layer across the surface of the wound, assisting with re-epithelialisation (Motamedi et al., 2021). Cell sprays have also been used in tandem with other skin substitutes, like skin grafts and biomaterial scaffolding (Motamedi et al., 2021).

Porous scaffolds seeded with cells provide structurally appropriate materials for more substantial wounds with deeper skin loss. For example, Dermagraft^®, a polyglactin mesh seeded with fibroblasts, which is approved for use in diabetic foot ulcers (Table 1) (Hart et al., 2012). Fibroblasts populating the matrix secrete their own ECM and incorporate with the wound below to increase the rate of healing (Hart et al., 2012). By producing multi-dimensional scaffolds, it is also possible to seed multiple cell types within layers to replicate the multi-layered structure of native skin. For example, Orcel^® is a collagen-based sponge seeded with keratinocytes in an upper gel layer and fibroblasts in a lower porous layer to replicate the epidermis and lower dermis (Table 1) (Still et al., 2003). This product is U.S. Food and Drug Administration-approved for the treatment of burn autograft sites and acute surgical wounds. Unfortunately, many of these engineered skin substitutes, both acellular and cellular, are not yet functioning as total skin replacements. In many cases, they still need to be applied in combination with a split thickness skin graft or acellular fillers (Halim et al., 2010; Wainwright and Bury, 2011). They also have a limited patient window, often only appropriate for wounds free of infection, with good vascularisation (Wainwright and Bury, 2011).

Beyond the biological level, there are other spaces for improvement, as the techniques used to produce engineered skin substitutes need to be scalable to the industry level (Dearman et al., 2021). Additionally, these products need to be financially accessible to patients. Many skin substitutes are associated with high cost (Table 1): OrCel^®, a bilayer dermal substitute, costs an estimated $27.78USD per cm², and Alloderm^® a decellularised human dermal matrix, costs an estimated $29.68USD per cm² (Mofid and Singh, 2009; Pourmoussa et al., 2016). High cost of skin substitutes remains a limitation not only to patient accessibility, but to scalability and manufacturing (Halim et al., 2010; Pangarkar et al., 2010). It is also becoming more important that tissue constructs are prepared using environmentally friendly and sustainable methods (Mahmud et al., 2023; Benko and Webster, 2023; D’Elía et al., 2023). The use of naturally-derived biomaterials with sustainable sources such as marine collagen and cellulose are compelling, but the extraction and production of these biomaterials remains a chemically-intensive process (Samavedi et al., 2014; El Knidri et al., 2018; Khan et al., 2022; Saji et al., 2022). The use of growth factors and signalling molecules to support cellular proliferation and differentiation within engineered constructs remains a resource-intensive and costly addition and alternative approaches to support cellular function are needed (Ren et al., 2020).

The electrical field strength, E, is one of the key factors determining the outcome following PEF treatment and is a culmination of the many independent parameters involved in PEF. This factor can be calculated as follows: Ε   kV / cm = v o l t a g e   d e l i v e r e d   U w i d t h   b e t w e e n   e l e c t r o d e s

This key factor, E, must be considered in relation to the desired outcome. As it is dependent on the width between electrodes, E will differ based on the PEF system and chamber used to deliver stimulation. As previously discussed, the transition between reversible electroporation and irreversible electroporation is threshold-dependent, based on the critical E reached (Edd et al., 2006; Lee et al., 2010). Above this critical threshold, cell viability is compromised, with permanent pore formation triggering apoptotic processes within the treated cells (Ganeson et al., 2018). Stronger electroporation at higher E can lead to enhanced extraction of intracellular compounds from the target cell.

The E applied to cell monolayers has been varied (Table 2). Electric field strengths of 0.3 kV/cm (pulse duration 5 m, frequency 1 Hz) and 0.8 kV/cm (pulse duration 100 µs, frequency 1 Hz) were cytotoxic to human dermal fibroblast populations in vitro (Gouarderes et al., 2020). A reduced E of 0.05 kV/cm (pulse duration 5 m) improved fibroblast migration in a wound scratch assay, but not to a statistically significant level (Gouarderes et al., 2020). Gouarderes et al did note that PEF-treated cells increased their expression of connective tissue growth factor (CTGF), VEGF-A, platelet-derived growth factor (PDGF)-A, and transforming growth factor (TGF)-α and reduced expression of TGF-β, and conditioned media from treated cells was able to stimulate the migration of unexposed cell populations (Gouarderes et al., 2020). The authors suggested that the enhanced expression of stimulatory factors by treated cells was due to release into the media during cellular necrosis, and cited mitochondrial stress as the cause of cell death rather than irreversible electroporation (Gouarderes et al., 2020). Das et al noted proliferative and migratory fibroblast responses after treatment of cells with 500 V (E not stated) (Das et al., 2020). In contrast to Gouarderes et al., Liu et al noted that an E of 0.003 kV/cm was not cytotoxic to mouse fibroblasts embedded in a collagen lattice (Liu et al., 2018).

Griffiths et al treated E. coli-contaminated collagen gels with PEF stimulation and successfully inactivated Escherichia coli at densities below 10³ CFU/mL (Griffiths et al., 2008) (Table 3). They noted that the effect was increased with increasing E (from 12 to 45 kV/cm). A subsequent study noted that treatment of the collagen gels with up to 60 kV/cm had no effect on collagen gel structure or the growth of osteoblasts on these scaffolds (Smith et al., 2009). PEF treatment of small microbial cells generally requires high E (10-14 kV/cm) to reach the threshold for irreversible electroporation (Bhat et al., 2019). In comparison, Giteru et al assessed the effect of PEF stimulation on the characteristics of a film composed of zein, chitosan, and poly (vinyl alcohol) (ZCP), focusing on phenotypic outputs (Table 3) (Giteru et al., 2020). Treatment of film-forming dispersions with electric field strengths of 0.9–3.4 kV/cm (pulse width 32 µs, frequency 10–220 Hz) produced different particle sizes, viscosities and loss moduli depending on E. Particle size increased with increasing E, while viscosity and loss modulus decreased. An optimal tensile strength was achieved using an E of 3.4 kV/cm and 50 Hz frequency (Giteru et al., 2020). In this case, Giteru et al (2020) achieved their desired biomaterial phenotype with a much lower E of 3.4 kV/cm and did not require the high E needed for sterilisation. When considering the development of skin and other tissue constructs, biomaterials must be both sterile and have the appropriate mechanical features. It therefore needs to be considered that E may need to change based on the stage of biomaterial preparation and that if these stages are being executed simultaneously, this may be difficult based on changing PEF requirements.

Modifying E may provide control over cell behaviour or viability, but this parameter differed largely in these experiments. More systematic testing is required to elucidate the exact effect that E may have on cells both in 2D cell culture and in 3D engineered tissues.

The shape of the pulses delivered by PEF can be exponentially decaying, sinusoid, or square, and can be monopolar or bipolar (Novickij et al., 2022). The choice of waveform can dictate the outcome following PEF, including whether electroporation is reversible or irreversible. Sinusoid and exponential pulses occupy time in the transient phases of the wave (i.e., increasing and decreasing), whereas square pulses rapidly reach and maintain their peak before rapidly decreasing. Given that PEF is threshold-dependent, square pulses are more effective at maintaining the electric field above the critical electrical field strength threshold compared to sinusoidal or exponential pulses. Therefore, square waves are much more effective at inducing irreversible electroporation compared to sinusoidal or exponential wave patterns (Novickij et al., 2022).

In addition to shape, the width of the pulse (τ_p) is also a variable that can affect PEF outcomes. Obtaining a square pulse shape is easier with longer pulse durations, and pulse durations below 50 ns may compromise the square shape of the pulse, which can defect to a sinusoidal shape (Novickij et al., 2022). To maintain a square pulse shape, thereby maximising the effectiveness of PEF treatment, pulse duration should be considered.

Compared to unipolar pulses, bipolar pulses are more efficient at inducing electroporation (Kotnik et al., 2001b). Bipolar square pulses also mitigate the electrolytic contamination due to release of metal ions from the electrodes during PEF (Kotnik et al., 2001a). Substantial electrolytic contamination was observed when using unipolar square pulses and this has been linked to decreased cell viability and function (Loomis-Husselbee et al., 1991; Stapulionis, 1999; Kotnik et al., 2001a). Therefore, when developing tissue constructs, care should be taken to select appropriate pulse shape and polarity to avoid unwanted cytotoxicity.

In the preparation of biomaterials with PEF, much of the literature neglected to comment on their choice of pulse shape, however those that did, most commonly reported the use of rectangular or square pulses (Griffiths et al., 2008; Smith et al., 2009; Sano et al., 2010; Giteru et al., 2020) (Table 3). These studies failed to comment on the polarity of the selected pulses. Use of PEF for cell treatment was much the same, with the common choice being square or rectangular pulses (Table 2) (Hess et al., 2012; Kumar et al., 2016; Nguyen et al., 2018; Gouarderes et al., 2020; Gouarderes et al., 2022), with the occasional comment on polarity (Fitzsimmons et al., 2008).

The number of pulses delivered (n _p), the pulse duration, and the frequency of pulse delivery can also affect PEF outcomes. Pulse number and duration impact the overall treatment time, thereby influencing the total energy input. Shorter pulse durations are generally preferred where cell viability needs to be conserved, while longer pulse durations aid the transfer of larger macromolecules across cell membranes (Rols and Teissié, 1998; Šatkauskas et al., 2012). The pulse frequency will affect the degree of pore formation and membrane permeabilisation (Lamberti et al., 2015; Novickij et al., 2022). Increased pulse frequency also augments the uptake of molecules by the targeted cells due to electrosensitisation, which should be considered in regards to the purpose of PEF treatment, for example, with drug delivery in electrochemotherapy (Bilska et al., 2000). This may be of assistance when combining PEF with delivery of pro-healing growth factors or drugs to cells or tissues (Berry-Kilgour et al., 2021). It has also been reported that pulse frequency influences the size of the pores formed during irreversible electroporation (Mi et al., 2019), which may influence the extraction of biological molecules from cells following PEF treatment.

Rahman et al noted that PEF-assisted electrohydrodynamic bioprinted sodium alginate droplets were altered by changing PEF parameters (Table 3) (Rahman et al., 2021). Using a modified continuous-style PEF delivery to bioink at pulse frequencies between 150 and 170 Hz, they reported that increasing frequency reduced the size and distance between droplets, but also reduced the consistency of droplet shape. Altering these parameters changed the printing outcome and should be considered based on the desired product.

Pulse number and frequency were also shown to influence decellularisation of structures (Table 3) (Phillips et al., 2010; Sano et al., 2010). Phillips et al applied 90 pulses at 1 or 4 Hz (E = 1.75 kV/cm) to rat carotid arteries in vivo by applying an electrode clamp to either side of the artery (Phillips et al., 2010). Five days after PEF treatment, they noted a loss of cells and decrease in nuclear staining in histological sections. Apoptotic cells resulting from irreversible electroporation were cleared by host immune responses to produce an acellular vessel. In the 1 Hz-treated vessels this effect was sustained beyond 7 days, but in the 4 Hz-treated vessels, repopulation of cells into the vessel was noted by day 7 (Phillips et al., 2010). Sano et al produced visible depigmented lesions on ex vivo porcine livers following PEF treatment with similar parameters (99 pulses, 0.25–4 Hz, E = 1 kV/cm, pulse width = 100 µs). Histological analysis of these lesions showed reduced cell number and preserved extracellular matrix structure, but no significant markers of decellularisation such as DNA content or DAPI staining were evaluated (Sano et al., 2010).

The effect of changing these PEF parameters on cells has not been extensively investigated. For the skin cell studies (Table 2), frequencies used were around 1 Hz. The effect of pulse number, duration and frequency on standardised cell outputs such as cytotoxicity or proliferation has rarely been reported in the literature. While standardised use of these outputs would aid comparison, functional assessments of cell behaviour have differed based on the cell type, for example, alignment of fibroblasts, or migration of keratinocytes. Liu et al found that increasing pulse frequency from 0.0002 up to 100 Hz increased fibroblast alignment in a collagen matrix (Liu et al., 2018). In another study, changing frequency from 0.1 to 1,000 Hz (E = 150 mV/mm, other parameters not stated) did not affect the migration capacity or speed of keratinocytes in vitro (Ren et al., 2019). Pulse duration also impacted fibrotic responses in cultured dermal skin substitutes (Gouarderes et al., 2022), where a long pulse protocol (10 pulses of 5 ms) increased MMP activity and reduced TGF-β1 expression to a greater extent than a short pulse protocol (8 pulses of 100 µs). Although, a transient decrease in collagen content was observed regardless of the protocol used (Gouarderes et al., 2022).

A highly conductive sample is at greater risk of arcing, a phenomenon where high E results in electrons jumping from one electrode to the other, rather than moving through the sample (Góngora-Nieto et al., 2003; Álvarez et al., 2006). This arcing decreases the efficacy of electroporation, increases sample temperature, and can cause damage to the electrodes (Barbosa-Cánovas et al., 1999; Álvarez et al., 2006; Oey et al., 2022). Because conductivity will determine the efficiency of PEF, the conductivity of chosen biomaterials needs to be considered. More conductive biomaterials (>4 m/cm) are showing promise in the engineering of electrically-responsive cardiac and muscle tissues (Gajendiran et al., 2017; Dong et al., 2020). Because high conductivity is linked to reduced efficiency, it may be easier to prevent arcing and loss of electroporation efficiency when incorporating PEF into the engineering of tissues such as epithelium or skin, which have lower conductivities (Saberi et al., 2019).

Temperature will increase following irreversible PEF treatment because of Joule heating (Davalos and Rubinsky, 2008; van Gemert et al., 2015). At higher temperatures, sample conductivity increases, which increases the likelihood of arcing (Barron and Ashton, 2005). In addition to considering success of PEF treatment, the desired outcome following PEF treatment should also be considered when optimising temperature. Increased temperature can enhance electroporation efficiency, but at the detriment of the target tissue, with a higher level of tissue damage and disintegration (Lebovka et al., 2005).

Whether or not the structure of the tissue construct must be maintained during and after PEF treatment should be evaluated when considering the sample temperature. For example, for the purpose of sterilisation or decellularisation of biomaterials, maintaining the appropriate structure is critical for success. Pre-sterilisation of temperature-sensitive collagen-based biomaterials were performed using cooling systems to reduce temperature and mitigate structural changes (Griffiths et al., 2008; Smith et al., 2009). Classical PEF studies have typically been conducted under chilled conditions or at room temperature (≤25°C) (Leong et al., 2014; Liu et al., 2017; Kantono et al., 2021; Karki et al., 2023). Complications may arise with PEF treatment of cell suspensions or 3D constructs, which must be maintained at physiological temperature. Liu et al looked at the effect of PEF on fibroblasts embedded within a collagen matrix at physiological temperature (37°C), but did not comment on the effect on the collagen matrix (Liu et al., 2018) (Table 2). It is well-known that collagen is temperature-sensitive (Miles and Bailey, 1999), and the increased sample temperature following PEF should be considered as it could affect the degradation rate not only of collagen, but of other physiologically relevant biomaterials. This becomes particularly important when looking ahead to PEF treatment of 3D tissue constructs and their stability in vivo or for prolonged culture periods at physiological temperature.