Novel, standardized pre-clinical surgical animal model to investigate the cellular and molecular mechanisms of ischemic flap healing

doi:10.21203/rs.3.rs-3058815/v1

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Method Article

Novel, standardized pre-clinical surgical animal model to investigate the cellular and molecular mechanisms of ischemic flap healing

https://doi.org/10.21203/rs.3.rs-3058815/v1

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published 16 Jan, 2024

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Background

Some of the most complex surgical interventions to treat trauma and cancer include the use of locoregional pedicled flaps and free autologous tissue transfer. While the techniques used for these reconstructive surgery procedures have improved over time, flap complications and even failure remain a significant clinical challenge. Animal models are useful in studying the pathophysiology of ischemic flaps, but when repeatability is a primary focus of a study, conventional in-vivo designs, where one randomized subset of animals serves as a treatment group while a second subset serves as a control, are at a disadvantage instigated by greater subject-to-subject variability.

Results

We present a novel, economical, and standardized pre-clinical animal model of excisional full-thickness wound closure and healing using axial pattern flaps that incorporates an anatomically named vascular pedicle. In this bilateral epigastric fasciocutaneous advancement flap (BEFAF) model, which simulates an actual clinical procedure of autologous tissue transfer, one flap heals under normal and a contralateral flap - under perturbed conditions or both flaps heal under conditions that vary by one within-subjects factor. Although we describe step-by-step methodological approach how to create such flaps as well as collect, process, and analyze flap survival data in larger size rodents, specifically, the rats (Rattus norvegicus), the BEFAF model is transferable and could be implemented in any mammalian species. We also stress the importance of taking and presenting high resolution histological stain images of the flaps that are indispensable for diagnostic result interpretation in the full-thickness wound healing studies.

Conclusions

BEFAFs may be used to investigate the spatiotemporal cellular and molecular responses to complex tissue injury and interventions simulating clinically relevant flap complications (e.g., secondary arterial, venous, or mixed ischemia) and therapeutic or surgical treatments (e.g., flap delay) in the presence or absence of confounding risk factors (e.g., substance abuse, irradiation, diabetes) or favorable wound-healing promoting activities (e.g., exercise). This technically challenging but feasible reconstructive surgery model eliminates inter-subject variability, while concomitantly minimizing the number of animals needed to achieve adequate statistical power. BEFAF also serves as an aid for teaching basic vascular microsurgery techniques that focus on precision, tremor management and magnification.

Excisional wound healing

bilateral flap

pedicled flap

fasciocutaneous flap

superficial inferior epigastric vessels

axial pattern flap survival

primary ischemia

reperfusion injury

secondary ischemia

autologous tissue transfer

According to the CDC’s National Center for Injury Prevention, each year trauma accounts for more than 39 million emergency department visits across the nation (1). Most of these cases require immediate or planned surgical intervention using compound flaps that are composed of different types of tissues such as full-thickness skin, muscle, bone or even cartilage. These flaps are further classified based on their (1) location (e.g., local, regional, distant); (2) arrangement of blood supply; (3) configuration and (4) the method of transfer onto the wound (soft tissue defect) (2, 3). For instance, a microvascular free flap is elevated from an uninjured donor site, fully detached, and transferred to the remote recipient site (Fig. 1A). Such flap is exposed to varying times of primary ischemia (PI) before being reconnected to recipient vessels by a technique known as anastomosis. By contrast, a local or regional pedicled flap remains attached at the harvest site by vascular subcutaneous tissue base known as a pedicle. During surgery, such a pedicled flap can be rotated a certain number of degrees around a pivotal point located at the base of the flap to reach and cover an adjacent, nearby, or distant recipient area (rotational, transposition or interpolated flaps) (Fig. 1B). In other cases, a pedicled flap is simply stretched toward and over a local or distant defect without altering the plane of the pedicle (advancement flaps) (Fig. 1C). Tissue deficits created in the donor site are closed primarily and generally heal relatively quickly and without complications.

Preclinical wound healing models distinguish among incisional and excisional wounds, flaps, and cutaneous grafts. A graft has no intact blood supply and therefore relies entirely on diffusion of nutrients and a number of new blood vessels derived from the recipient wound bed. By contrast, a flap contains native blood supply. In an axial pattern flap (APF) the blood is delivered via a dominant and well-defined cutaneous artery that is often accompanied by a large-scale vein. In a random pattern flap (RPF), the blood is supplied via an unnamed subdermal perforator plexus. Due to their increased length and more robust inherent blood supply, APFs are ideally suited for reconstruction of large and deep soft tissue defects that are beyond the reach of shorter and less predictable RPFs. Nevertheless, the survival pattern of APFs is typically based on the integrity and extent of their main feeding and draining vessel(s). Thus, a distal portion (the very tip) of the APF that exceeds the distance of the underlying longitudinal artery may behave as a graft or an RPF: after a reconstructive surgery, these distal tissues stay deprived of oxygen (ischemic) significantly longer depending on the rate and extent of neovascularization. To a certain degree, naturally occurring hypoxia (the lack of oxygen) will stimulate the APF-infiltrated and APF-residing cells to release pro-inflammatory cytokines and angiogenic factors contributing to the inosculation (connection of pre-existing vessels to the vessels of recipient wound bed and sidewalls) and angiogenesis (the ingrowth of new vessels). But if hypoxia persists, sustained inflammation combined with local accumulation of reactive oxygen species (ROS) and lactic acid buildup will lower the intracellular pH and cause severe DNA damage. As a result of extensive apoptosis-related protein activation, depleted bioenergetic resources such as NAD + would trigger massive cell death and, subsequently, devitalized tissue breakdown (necrosis) (4–6).

Despite recent advances in surgical techniques and post-operative monitoring that contribute to a high (~ 85–95%) flap success rate (7–10), complications associated with vascular compromise and ischemic partial- or full-thickness necrosis remain a significant clinical problem, particularly in angiopathic, obese, post-irradiated, actively smoking and/or alcohol-abusing individuals (11–20). In addition to inducing debilitating patient stress, flap failures inordinately contribute to health care costs due to prolonged hospitalization, readmission and revision surgery (21). The success rate of salvaged flaps ranges from 28% to over 90%, depending on the type of the flap being used, the etiology of its failure, the timing of its salvage and the overall experience of medical center surgeons and staff (22–24).

Experimental translational plastic and reconstructive surgery research animal models of complex full-thickness wound closure and healing using RPFs and AFPs are indispensable tools in studying physiological and pathophysiological responses to acute intermittent hypoxia and ischemic tissue injury (25). These models provide insights into a spatiotemporally regulated activation machinery of integrated signaling pathways that induce transcription of genes controlling the survival, motility, proliferation, and differentiation of multiple functionally distinct cell populations. Under the influence of diverse endogenous stimuli these locally residing, or newly arrived wound-bed cells are capable of either maintaining or ceasing inflammation. Some of them promote vascularization through angiogenesis and lymphangiogenesis, change hemodynamics and induce tissue remodeling. A complete understanding of each physical and biological facet of the reparative ladder resulting in a fully recovered flap and completely closed wound is essential to predicting the clinical outcomes of the reconstructive surgery.

Many preclinical research animal wound healing models are primarily designed to investigate tissue ability to progress through different phases/stages of wound healing under unperturbed or minimally perturbed (control/sham) and experimental “treatment” conditions. The latter includes irradiation (26), photodynamic therapy (27), treatment with pharmacological agents (28–30), recombinant peptides (31), plasmid DNA or siRNA compounds (32), synthetic or natural products (33, 34)) etc. Sometimes the investigators aim to mimic severe medical complications (e.g., vascular thrombosis, edema, ulcerative and infected wounds) (35) or reproduce well-known comorbidities (e.g., obesity, diabetes, ageing, smoking, alcohol intake etc.) (36–38). Different routes and/or regimen of single or combined drug administration (39, 40), ischemic preconditioning (41, 42), delaying (43) and numerous perioperative flap treatment or handling strategies (44–48) have been evaluated for their efficacy at reducing the risk of compound flap loss. However, in all conventional experimental studies one randomized animal group typically serves as a control, while another group undergoes the desired treatment(s). The results obtained from such studies suffer from reduced statistical power due to high subject-to-subject variability. Pre-surgical fluctuations in baseline levels of activated proteins/genes and circulating factors may mask the moderate and less robust effects of chosen treatment(s), especially if it is applied directly onto the flap. Thus, to achieve stronger statistical power, a larger number of animals must be used.

Here we introduce a novel, standardized complex full-thickness wound closure model, which is based on a bilateral flap. In this versatile, scalable, transferable, and reproducible model, a treated or perturbed ventral (APF) or dorsal (RPF) flap is paired with a contralateral control ventral or dorsal flap on the same animal which heals under similar physiological or pathological conditions. The APFs may be rotated along their axis to end up on the dorsal side and vice versa: the caudally created RPFs may be transposed towards a ventral side. Significantly greater control for between-animal variability is obtained while sacrificing fewer animals.

4.1. Overview of BEFAF model

In the current adaptation of the APF model to reconstruct an abdominal wound, we employ a bilateral epigastric fasciocutaneous advancement flap (BEFAF), which is based on two ventral APFs supplied by the caudal superficial inferior epigastric artery (SIEA) and drained by superficial inferior epigastric vein (SIEV). SIEA and SIEV branch from the femoral artery/vein in the left and right groin regions. In clinical practice, SIEA-based epigastric flaps are used either as unipedicled or bipedicled advancement or transposition flaps to cover ipsilateral or contralateral wounds on the abdomen, perineum or hindlimbs, or as free flaps to repair facial or distant breast defects as a result of trauma, burns, tumor resection or infection (49–57). First, we create a full-thickness soft tissue defect in the upper abdomen region: a piece of fasciocutaneous tissue measuring between 0.5×3 cm and 1×3cm is excised from the top portion of a flap prior to its elevation. Due to the absence of extensive physical manipulation, the excised tissue thus reflects the basal activation, expression and/or spatial distribution patterns of genes, microRNAs, and proteins. In routine histological tissue sample analysis, it represents morphologically normal unwounded skin (control). The qualitative features as well as quantitative measures obtained at this baseline (0 h) can be readily compared to the ones obtained within the proximal (P), middle (M) and distal (D) flap segments at specific time points following an excisional and ischemic injury. Taking a second biopsy from the D segment of the raised flap prior to its advancement and final wound closure results in a fully advanced APF with a clinically recommended 3:1 or 2.5:1 height to width ratio. If the combined excised tissue area is insufficient for analysis (less than 1.5×3cm), then a third biopsy may be acquired at any intermediate time point via minor survival surgery. The degree of tissue necrosis and overall survival, expressed as flap viability percentage (FV%), is evaluated by digital planimetric analysis using digital photographs of ready-to-be harvested flaps at any designated postoperative day (POD) (46). These endpoints typically represent the peaks of major wound healing stages: inflammation (POD1 to POD3), proliferation (POD 5 to POD 7) and short-term or long-term remodeling (POD 9 to 14 and beyond). The rate of flap healing/failure may be calculated through this sequential planimetric analysis. The most common variables for comparative BEFAF analysis are listed in Table 1.

Importantly, with this BEFAF model, one may fully recreate microvascular free flap surgery. One or both flaps can be entirely detached from their primary blood supply followed by end-to-patch anastomosis of smaller caliber vessels ranging from 0.3 mm to 0.8 mm in diameter. This technique is feasible in rats but requires using a microvascular surgery microscope or high magnification loupes (58–63). To simplify this highly specialized procedure and make it more readily accessible to non-medical researchers, we mimic anastomosis by using atraumatic vascular clamps. The clamps are gently applied onto both feeding and outflow vessels in one or both flaps for as long as it would presumably take the surgeon to separate the tissue, transfer and reconnect it to the exposed vessels of the wounded area. The global PI typically lasts no more than 2 hours, but depending on the experimental aims, flap type and animal species used, the induced PI time may be tailored to be either shorter or longer. Furthermore, the BEFAF model presents an opportunity to examine the effects of acute or late secondary ischemia (SI). SI may be artificially induced by temporary closure of the SIEA, SIEV or both vessels after a certain primary reperfusion (REP₁) period following global PI.

Table 1

Suggested pre-operative or perioperative flap handling or treatment conditions for the comparative short-, intermediate- or long-term whole flap or segmental gene or protein expression, cell signaling, extracellular vesicle release, neovascularization, tissue survival and other analyses within one subject.Abbreviations: PI - primary ischemia; SI - secondary ischemia, AO – arterial occlusion, VO – venous occlusion; AVO – arteriovenous occlusion, REP₁ – primary reperfusion.
Left flap	Right flap
Control naïve flap (raised and placed down)	Ischemic flap (e.g., 2 h of PI)
Control naïve flap (raised and placed down)	Flap with inserted sterile silicone sheet
Control naïve flap (raised and placed down) APF	Permanently ligated artery, vein, or both^# RPF
Global PI (e.g., 2 h)	Global PI (e.g., 6 hours)
Global PI (e.g., 2 h) REP₁ (e.g., 4 h) Global SI (e.g., 2 h)	Global PI (e.g., 2 h) REP₁ (e.g., 12 h) Global SI (e.g., 2 h)
Global PI (e.g., 4 h) REP₁ (e.g., 12 h) Global SI by AVO (e.g., 2 h)	Global PI (e.g., 4 h) REP₁ (e.g., 12 h) Global SI by AVO (e.g., 6 h)
Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by AO (e.g., 2 h)	Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by VO (e.g., 2 h)
Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by AO (e.g., 2 h)	Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by AO (e.g., 6 h)
Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by VO (e.g., 1 h)	Global PI (e.g., 2 h) REP₁ (e.g., 6 h) SI by VO (e.g., 3 h)
Ischemic preconditioning (e.g., 10 min × 3 times)	No ischemic preconditioning Control naïve flap (raised and placed down)
Delayed flap (e.g., 7 days)	No delay
Implantation of control PVA sponges	Implantation of drug-preloaded PVA sponges
Saline injection	Drug injection
Intradermal delivery of non-targeting control siRNA, shRNA, or antisense oligonucleotide	Intradermal delivery of targeting siRNA, shRNA, or antisense oligonucleotide
Topical treatment with control vehicle	Topical treatment with an agent of interest
No anastomosis applied	Anastomosis

# - flap ligation is a positive control used to estimate the maximal extent of flap failure upon complete blockade of blood flow through the main artery. This procedure turns the APF into RPF.

Such experimental design directly mimics arterial (AO), venous (VO) or mixed arterio-venous (AVO) occlusion or thrombosis. These types of vascular complications most frequently occur within first 72 hours after free tissue transfer and are a major contributor to flap loss (64–67). In the BEFAF model, flap relief from the SI would simulate a clinical flap re-exploration and salvage procedure. Once the arterial, venous, or arteriovenous congestion is alleviated, the flap would go through the secondary reperfusion (REP₂) and recovery period.

Both REP₁ and REP₂ are else known as reoxygenation events during which blood supply returns to the ischemic regions of the tissue. Depending on the degree of flap hypoxia and the rate of blood return, the restoration of circulation may result in mild or severe inflammation and oxidative damage through the induction of oxidative stress rather than (or along with) restoration of normal tissue function. This phenomenon of ischemia-reperfusion injury (IRI) is well-known and has been widely explored mechanistically (68, 69). The BEFAF model allows investigators to assess the tolerance of male vs female or treated vs untreated animal flaps to the selective PI and/or SI events, generating probit analysis curves and calculating the critical ischemic time (CIT₅₀) values when 50% of flaps become totally necrotic (Table 2) (28).

One additional advantage of using the BEFAF model is that during an inflammatory phase of wound healing such flap produces copious amounts of wound exudate (WE). This growth factor-, cytokine-, chemokine-, metalloproteinase- and extracellular vesicle (EV)-enriched fluid freely travels from the caudal to the cranial regions of ventral flaps facilitating the diffusion of vital healing factors that in turn promote cell migration across the wound bed, cell proliferation, metabolism and aid the autolysis of damaged tissues (70). The flap can be pre-opened for the manual collection of WE into a sterile 1000 µL pipette tip after a flap is gently squeezed at the base. But more conveniently, one or more sterile polyvinyl alcohol (PVA) sponges can be implanted underneath the flap to absorb the WE over time (71). Control sponges may be subcutaneously placed in the midline between the left and right flap or outside of each flap further away from the wound. The sponges are eventually removed by opening the pockets in the respective portion(s) of the flap(s) through minor survival and/or non-survival (endpoint) surgery. Alternatively, the PVA implants may be preloaded or continuously injected with a control vehicle or an experimental treatment solution to evaluate its impact on the migratory properties of different wound-bed cell populations, rate of granulation tissue (GT) formation and its health (thickness, vascularity), lymphangiogensis as well as deposition of extracellular matrix (ECM) and collagen fibers (72–75).

Table 2

Experimental research animal model designs for estimating the critical primary and secondary ischemia times for single-pedicled flaps in male and female subjects at 95% power and planned minimum 3 h time gap between two anesthesia events on the same subject.
Animal number per group excluding attrition		Left flap	Right flap
Study Design A
Group A	Group B
7 female	7 male	No PI	Ligation^#
7 female	7 male	PI for 2 h	PI for 8 h
7 female	7 male	PI for 4 h	PI for 10 h
7 female	7 male	PI for 6 h	PI for 12 h
N = 28	N = 28	n = 7, r = 1 per group	n = 7, r = 1 per group
Study Design B
7 female	7 male	No PI	PI for 2 h
7 female	7 male	PI for 2 h, REP for 6 h, SI for 1 h	PI for 2 h, REP for 6 h, SI for 4 h
7 female	7 male	PI for 2 h, REP for 6 h, SI for 2 h	PI for 2 h, REP for 6 h, SI for 6 h
N = 21	N = 21	n = 7, r = 1 per group	n = 7, r = 1 per group
Study Design C
7 female	7 male	No PI	PI for 2 h
7 female	7 male	PI for 2 h, REP for 12 h, SI for 1 h	PI for 2 h, REP for 12 h, SI for 5 h
7 female	7 male	PI for 2 h, REP for 12 h, SI for 3 h	PI for 2 h, REP for 12 h, SI for 7 h
N = 21	N = 21	n = 7, r = 1 per group	n = 7, r = 1 per group
Study Design D
7 female	7 male	No PI	PI for 2 h
7 female	7 male	PI for 2 h, REP for 24 h, SI for 2 h	PI for 2 h, REP for 24 h, SI for 8 h
7 female	7 male	PI for 2 h, REP for 24 h, SI for 4 h	PI for 2 h, REP for 24 h, SI for 10 h
7 female	7 male	PI for 2 h, REP for 24 h, SI for 6 h	PI for 2 h, REP for 24 h, SI for 12 h
N = 28	N = 28	n = 7, r = 1 per group	n = 7, r = 1 per group

# - vessel ligation is a positive control used to estimate the maximal extent of flap failure upon complete blockade of blood flow through the main artery.

Although there are many different extensions of the BEFAF model discussed in the section below, in this publication we provide step-by-step instructions on the creation of BEFAFs for the intermittent collection of tissue biopsies and a final evaluation of flap survival at one designated endpoint after the surgery in rats (Rattus norvegicus), because work with live rats is considered the gold standard in the current microsurgical training (76). The chosen animals may be any desired strain, sex, age, and weight – due to the differences in skin anatomy and ischemic tolerance (female vs male), or wound healing efficacy (young vs old) - they all come with different advantages. Experimental animals may be housed under routine or specialized husbandry conditions and fed ad libitum or a specialized (e.g., alcohol-containing) diet. They may be randomly selected and divided into two or more experimental groups receiving any control or experimental treatment prior to flap raising surgery.

In our example of BEFAF surgery workflow provided below, the left-side flap is subjected to 4 h of global PI, 2 h of REP₁ and 4 h SI by VO, while right-side flap is subjected to 4 h of global PI, 2 h of REP1 and 4 h of SI by AO. Thus, in principle, the presented method is a fusion of three clinically relevant experimental techniques (pedicled flap advancement, free microvascular flap surgery and flap salvage). We show representative photos of experimental male or female animals. As a visual aid, we used black non-absorbable silk sutures in every but the final suturing step. These sutures must be replaced with non-absorbable sutures where and when indicated in the protocol. Finally, we discuss data collection and analysis strategies showing representative micrographs of flap tissue sections that were obtained from the animals subjected to identical or similar procedures with varying REP₁ time, SI types and data collection endpoints.

4.2 Experimental animal preparation for survival surgery

To reduce the time of animal exposure to anesthesia during the major survival surgery, the animals should be shaved 16–24 h prior to flap raising procedure:

The heating pad is set for 122°F (50°C) and covered with two surgical underpads.
The anesthesia machines is turned on to prefill the padded rodent sedation chamber with 4–5% induction isoflurane gas.
The animal is placed in the chamber and kept there for up to 4 minutes.
3% isoflurane flow is then redirected into the nose cone attached to the anesthesia maschine.
The sedated rat is removed from the chamber and placed onto the heating pad in a prone position.
The lubricate ointment is generously applied onto animal’s eyes.
The rat is placed back into a supine position and its fur is shaved until the nude skin is exposed.
The top surgical pad containing shaved fur is removed.
Shaved skin areas are wiped with betadine solution, which is copiously applied onto a non-woven sponge.
Animal weight is recorded in the post-operative animal monitoring sheet (POAMS) (see Additional File 1).
The animal is returned to the cage and monitored until awakened.

4.3 Major survival surgery

4.3.1 Preparation

Appropriate PPE is donned by the surgeon.
The HDPE tubes, 10% formalin-prefilled containers and histology cassettes are labeled for appropriate specimen.
The vascular clamps, that are being stored in enzymatic solution, are removed, washed with distilled water and then placed in an Eppendorf tube containing 70% alcohol.
The hot glass bead sterilizer is turned on to preheat.
The heating pad is set to reach 122°F (50°C). It is covered with four surgical underpads: 1 extra-large and 3 medium or small.
If an optional PVA sponge implantation is desired, then two sterile 9 mm x 15 mm ear wicks are placed onto 100x20 mm tissue culture dish prefilled with sterile PBS. These wicks are then cut into three piece per each using a disposable scalpel. The lid of tissue dish is closed to prevent the PVA sponges from drying.
The dose of analgesic agent is prepared for each individual animal based on the recorded animal weight (for example, our rats received 1.0-1.2 mg/kg of Buprenorphine-HCl-SR formulation subcutaneously (sc)).
Dewar flask is prefilled with liquid nitrogen (LN₂).
Isoflurane and oxygen tank levels are checked, and the proper amount of isoflurane is added to the gas anesthesia machine chamber before the experiment if needed. The anesthesia machine is turned one to prefill the padded rat sedation chamber with 4–5% isoflurane gas.

4.3.2 Flap design

To prevent tension-associated flap failure, the BEFAF model allows taking only a limited number of tissue biopsies from the distal clinically most relevant part of the flap prone to failure. Additional biopsies, if required, may be obtained from the regions that lie along and close to the suture line, however, such biopsies introduce an additional variable. Therefore, in flap survival studies, no interference other than collection of biopsies from the distal portion of the flap throughout the entire course of experiment is strongly recommended. For a flap with initial measurements of 9.5×2.5 cm or 9.5×3 cm, the maximal flap advancement is 2 cm, which leaves a healing area of 3:1 or 2.5:1 length to width ratio. Consequently, one can obtain two 1 cm or four 0.5 cm wide biopsies from each flap. The 1×3 cm tissue strip is big enough to be divided into three or four segments for postoperative imaging or biochemical analysis, especially if sample pooling is allowed. On the other hand, such long and wide initial flap can be created only in adult, over 500 g rats. These are usually males. Female rats rarely exceed 350 g of weight. Therefore, for smaller animals, flap dimensions should be downsized appropriately, keeping in mind that a healing flap shrinks approximately 0.5–1 cm, which may partially limit animal movements and cause distress.

The animal is placed into the anesthetic sedation chamber and kept there for up to 4 minutes.
3% isoflurane flow is redirected into the nose cone attached to the anesthesia machine.
The sedated rat is removed from the chamber and placed onto the heating pad in a prone position.
Animal’s eyes are lubricated by applying the ointment, and analgesia is injected sc.
The animal is turned back to the supine position. The use of medical tape for animal stabilization during the flap design procedure is not recommended, as the animal skin should be in the most natural and relaxed condition.
Flap boundaries are identified by applying a pre-cut cardboard 9.5×3 cm flap design template and drawing a single dotted line with a skin marker. After removing the template, a second dotted line is drawn above the first just a few millimeters apart. Finally, a solid line is drawn through the inner dots as shown in Fig. 2A.
Flap areas that will be used for the excisional biopsies are then marked. In the example provided herein we defined four 1×3 cm biopsy areas:

A) B₀ – baseline.

B) B_PI – to be excised after the end of global primary ischemia (PI) and prior to reperfusion.

C) B_VO – to be excised after the end of secondary venous ischemia.

D) B_AO – to be excised after the end of secondary arterial ischemia (Fig. 2A).

Assuming the morphology and cellular activities in left and right side are similar under identical conditions, we aim to take B₀ and B_PI biopsy only from one of the flaps. Note: in an alternative experimental design, one can excise four 1×1.5cm biopsies out of each flap that would include in a dedicated B_0, B_PI and B_VO or B_AO sample plus one extra sample, e.g., B_REP1 - to be taken after the end of primary reperfusion prior to SI. However, the created gap resulting from partial tissue excision would need to be protected by gauze during intermediate incubation periods prior to final wound closure (see Additional File 2, Fig. S1).

4.3.3 Flap raising

An incision on the top of the flap is made using straight Iris Supercut scissors. The cut is deepdend. A portion of biopsy area is released from the fascia using Mayo dissecting scissors (Fig. 2B).
The baseline (B₀) biopsy area is lifted using Adson tissue forceps. This portion of skin is then cut out using Metzenbaum-Nelson scissors (Fig. 2C, left panel).
For HIS/IHC, the excised specimen is divided using a straight Iris Supercut scissors. The specimen is placed into a pre-labeled histology cassette, which is submerged in 10% formalin. For long-term tissue storage, the remaining specimen pieces are placed into the pre-labeled HDPE tube, which is sealed and submerged in LN₂.
After a re-entry to the existing pocket, the tissue is dissected underneath the flap by slowly moving Mayo scissors towards the pedicle (Fig. 2C, right panel). It is important not to open the scissors too wide beyond the medial boundaries of the designed flap.
Straight Iris Supercut or Metzenbaum-Nelson scissors are used to cut through the inner sidewall of the flap up to its end (Fig. 2D, left panel). The same is done for the opposite side (Fig. 2D, right panel). Some portions of the flap may still be attached to the underlying muscle by vascularized fascia (Fig. 3A, left panel). If so, a flap is lifted, and the fascia is cauterized so that the flap can be freed up to its base (Fig. 3A, right panel).
Straight Iris Supercut or Metzenbaum-Nelson scissors are used to cut through the lateral solid line of the flap roughly to its midline (Fig. 3C, left panel). From this point forward only the superficial layer of the flap should be cut through to avoid the damage of lateral branch of the SIEA (Fig. 3C, right panel). The same is done for the opposite side.
The cautery device is used to cut through the tissue of the elevated flap until its base (Fig. 3D). The same is done for the opposite side.
Both flaps are returned into their original horizontal position.

4.3.4 Physical vessel separation and selective clamping to induce primary ischemia (PI)

The pedicle is exposed. Some rats have abundant fatty tissue obscuring a view to the emerging SIEA/SIEV bundle from femoral artery/vein. Such cases require a deeper dissection and removal of more fascia around the thigh and the vessels (Fig. 4A). Once the vessels appear clean, each flap is wrapped in sterile gauze moistened with sterile DPBS to preserve them from drying and return to the original horizontal position (Fig. 4B). Note: to minimize variation, the temperature of DPBS should be kept constant unless the objective is to compare the viability of flaps exposed to warm or cold ischemia.
Isoflurane flow should be decreased to 2%.
Holding the tip and the center portions of the wrapped flap, a flap is positioned over the animal’s thigh so that the pedicle appears in an extended position (Fig. 4C).
The SIEA and SIEV vessels are irrigated in 1% Lidocaine HCl solution applied dropwise.
At this point the vessels of the pedicle may be examined more closely. The most suitable site for physical SIEV or SIEA separation is based on vessel anatomy and the surgeon’s left- or right-handedness. Vessel anatomy in male and female rats slightly differs. In contrast to females (Fig. 4D, left panel), male SIEA may be less pronounced and run very close to SIEV without making large loops (Fig. 4D, right panel), which may pose a challenge for their separation without an aid of magnifying spectacles and LED headlight.
For SIEV isolation the Halsey micro needle holder scissors are used. Once the largest gap between the SIEA and SIEV before the point of SIEV bifurcation is identified (Fig. 5A), a 4 − 0 size needle with attached silk suture is passed between the SIEA and SIEV in a direction shown in Fig. 5B. When the needle shows on the opposite side of SIEV, the scissors are released. The needle is grabbed close to the tip and slowly pulled upwards until the suture is underneath the SIEV (Fig. 5C). CRITICAL: if one of the vessels gets accidentally punctured, the bleeding can be stopped by clamping, raising both vessels and ligating them with LigaClip tool.

When properly ligated and cut in half, vessels should not bleed (see Additional File 2, Fig. S2A and Fig. S2B). A ligated flap may serve as a negative control for BEFAF survival. The flap (right or left) that has been ligated and the time of ligation should be immediately marked in the POAMS “Surgery notes” field (Additional File 1).
The suture is cut, and a loose knot is tied around the isolated vessel (Fig. 5D). Note: The 4 − 0 size needle should make a passage wide enough for a subsequent application of vascular clamp. It is possible to carefully widen this gap by using microscissors and lifting the attached suture loop, but it may increase the risk of vessel damage.
Similarly, the SIEA is separated on the opposite flap (Fig. 5E). If the SIEA forms a significant branch, a suture should pass right before the bifurcation point as shown in Additional File 2, Fig. S2C.
In the POAMS form (Additional File 1) the “Surgery Notes” field is used to indicate which vessel has been isolated by looping on the left and the right side of the animal.
Slightly or fully curved smooth jaws micro forceps are used to lift fascia. Then Iris microscissors are used to make a small tunnel below the tied loop on the top and bottom of the pedicle (Fig. 6A). This passage is widened by inserting the microscissors underneath the pedicle and slowly opening them to dissect the remaining tissue (Fig. 6B). This step is repeated for the opposite side (Fig. 6C).
Vascular clamp-applying forceps are used to insert one B-2V vascular clamp onto the pedicle with an already separated SIEV (Fig. 6D). Using slightly curved microforceps, the flow below an applied clamp is gently pushed towards the femoral vein to make sure that the blood outflow from the vein is fully ceased. It is normal to see some backflow from the femoral vein.

Note

If the venous blood surpasses the clamp, there may be too much fascia left around the vessel. If a problem persists after repositioning of a clamp, a second (smaller) clamp may be applied making sure there is no blood entrapped between the two clamps (see Additional File 2, Fig. S2D).
Once the clamp is in a proper position, a countdown timer is immediately set and started for the desired duration of the PI (e.g., 4 hours).
Steps 11 and 12 are repeated for an opposite side. In the case of arterial occlusion, the curved microforceps should be used gently to push blood away from the site of the clamp going distally towards the flap. If the clamp is adequately applied, no influx of blood will be seen in the artery between the clamp and the curved microforceps.
The flap is held vertically by regular forceps, and the microscissors or fine-tip cautery is used to widen the passage by dissecting tissue along the clamped SIEA/SIEV bundle (Fig. 6E). The angle between the isolated clamped pedicle and the flap should be ~ 45–60°. The same step is repeated for the opposite side.
Both flaps are returned to their original horizontal position. The moisture level of the gauzes that are wrapped around each flap is checked. If needed, an additional amount of sterile DPBS is applied. If the underpad is wet, it should be replaced to avoid hypothermia of the animal.
At this point the flap is supplied by residual microvascular plexus. To remove this feed the entire base portion of the flap is cut superficially using the Iris Supercut scissors (Fig. 7A). Note: this step can be omitted for local/regional pedicled advancement or rotational flaps, since these do not simulate a free flap situation.
The tissue at the base of the flap is cauterized while holding the flap vertically (Fig. 7B). At this point, the flap becomes connected to the animal only by the clamped pedicle (Fig. 7C). CRITICAL: Without pulling the pedicle (it may cause undue strain on the vessels), the flap is returned into its original horizontal position (Fig. 7D), and this step is repeated for the opposite side.

4.3.5 Partial flap closure

The 2% isoflurane flow should be decreased to 1%.
The flaps are sutured to the wound boundaries by using buried simple-interrupted suturing technique and non-absorbable silk suture (Additional File 2, Fig. S3) in the order and directions as suggested in Additional File 2, Fig. S4.
First, the bottom corners of a longer flap (which did not undergo biopsy excision) are sutured, and the same procedure is performed on a shorter flap (Additional File 2, Fig. S5). It is important to begin by suturing the base of the flap in order to minimize strain on the vascular pedicle.
Second, the upper corners of a longer flap are sutured followed by the center (Fig. 7E, left panel) and the remaining suturing points at the gaps (Fig. 7E, middle panel). A water-tight closure of the top and bottom parts of the flap is not necessary, but enough to prevent exposure of the clamp or vessel isolation loop and excessive accumulation of blood onto the future biopsies (Fig. 7E, right panel).
On the opposite side, the upper corners of a shorter flap are sutured using a non-absorbable silk suture. Finally, the center and the remaining points are sutured using an absorbable suture. Note: the guiding dots for suturing of this flap will not exactly match since the flap will be slightly advanced (Fig. 7F).
Isoflurane flow is decreased to 0% so that the animal would receive 100% oxygen. When it starts showing the signs of awakening on the operating table, it is returned to a cage to recover.
The instruments are cleaned from blood and dry-sterilized using the hot bead Germinator.
All waste is removed, and sharps are disposed in specified containers.
The cage is kept in the vicinity to monitor the animal’s well-being and activities more closely. If it starts extensively picking on the sutures, the Elizabethan collar should be applied.

4.3.6 Primary reperfusion (REP₁)

Fifteen minutes prior to the planned end of PI, the animal is placed into the anesthetic chamber for up to 4 minutes.
Meanwhile, two new underpads are placed onto a heating pad.
3% isoflurane flow is redirected into the nose cone.
The sedated animal is removed from the chamber and placed onto the heating pad in a prone position. The lubricate ointment is applied onto both eyes prior to turning the animal over in a supine position.
Isoflurane flow is adjusted to the maintaining 2% (for larger animal) or 1% (for smaller animal).
The flaps are gently wiped with DPBS-moistened gauze pad to remove any aggregated blood. Then the top underpad can be removed.
The degree of ischemia is inspected visually in both flaps. After 2 h of PI, flap color should be significantly darker than the surrounding skin, and even more so after 4 h of PI (Fig. 8A). Note: if the color does not significantly differ, it may indicate that the vascular clamping was insufficient or ineffective. With the extent of usage, the gripping power of reusable clamps gets weaker, and caution must be taken to prevent the clamp from falling off the pedicle in an active animal.
The suture is disrupted at points #4, #5, #6, #17 and #18 from the top portion of the longer (in this example, left) flap, followed by excision of the B_PI biopsy (Fig. 8B).
The sutures are removed from points #1 and # 9 (also #3 if needed more space) at the lower midline and an inner corner of the flap base on both sides (Fig. 8C, left panel).
Left flap is gently lifted (Fig. 8C, right panel) to remove the clamp by gripping it with the vascular clamp-applying forceps.
The pedicle is observed for restored blood flow (Fig. 8D). Note: we take note whether, upon clamp removal, there is some movement of venous blood. Small thrombi, if any, can be removed by applying a gentle sliding force with non-serrated microforceps. The vessels can be irrigated with 1% Lidocaine HCl solution applied dropwise after touching the pedicle.
Once the blood shows a sign of flow within the pedicle, the countdown timer is set and started for the desired primary reperfusion (REP₁) interval (e. g. 2 h).
Steps 9 to 13 are repeated for the right flap. Then, flap corners are sutured back to their original position using silk suture.
The remaining sutures on the left flap are disrupted. The top portion is reconnected to the tissue using silk suture. IMPORTANT: Then, an absorbable suture is used to connect the center and remaining parts so that both flaps are advanced equally.
Steps 6 to 9 from Section 3.5 are repeated.

4.3.7 Secondary ischemia (SI)

Fifteen minutes prior to the planned end of REP₁, the animal is anesthetized and prepared for the procedure as described previously.
3% (for larger animal) or 2% (for smaller animal) isoflurane flow is redirected to the nose cone.
The appearance of both flaps is visually inspected to assess the approximate percent of restored blood flow. Figure 9A shows a case of efficient (~ 73%) reperfusion of a flap only 2 hours after a designated 4 h PI time, which is well above the expected average (77).
The suturing points #2, #3 and #10 are disrupted at the lower sidewall and outer corners of flap base on both sides (Fig. 9B).
Left flap is gently lifted to inspect whether the blood flow is fully restored in the pedicle (Fig. 9C). Caution: some animals may develop spontaneous arterial or venous or mixed thrombosis after the PI (Additional File 2, Fig. S6). If a drop of heparin and 1% lidocaine does not resolve the thrombus and if the flow is not restored after a gentle push toward the groin area, the flap will be lost and should be excluded from the study. In such case, an opposite flap may be used as 4 h PI data point or any selected SI data point without a matching pair.
B-1V vascular clamp is applied onto an isolated SIEV (Fig. 9D). While applying the clamp, the suture that separates both vessels is removed. The timer is set and started for desired venous occlusion (VO) period (e. g. 4 h).
B-2V vascular clamp is immediately applied onto an isolated SIEA on the opposite side. The timer is set and started for desired arterial occlusion (AO) period (e. g. 4 h).
The bottom of both flaps is then closed using silk sutures.
Steps 6 to 9 from Section 4.3.5 are repeated.

4.3.8 Secondary reperfusion (REP₂)

Fifteen minutes prior to the planned end of AO/VO, the animal is anesthetized and prepared for the procedure as described previously.
2% (for larger animal) or 1% (for smaller animal) isoflurane flow is redirected to the nose cone.
The appearance of the flaps is examined to evaluate the efficacy of SI. VO flap should look significantly darker than AO flap (Fig. 9E). VO flap may be thickened by accumulated blood, which may form dark aggregates around the edges of the flap. These aggregates can be removed by flap irrigation with DPBS.
The sutures from the top portions of both left and right flaps are disrupted to excise the B_VO and B_AO biopsies, respectively (Fig. 9F).
The entire bottom of both flaps is opened widely to remove the clamps. The time of clamp removal is immediately indicated in the POAMS form to track the timing of secondary reperfusion (REP₂).
The pedicles are observed for the restored flow. The thrombi are removed if needed, and both vessels are irrigated in 1% Lidocaine HCl applied dropwise.

4.3.9 Full flap closure

The bottom of both flaps is sutured using an absorbable suture.
The top of naïve wound bed tissue is narrowed by suturing the upper corners together (Fig. 9G, left panel). The top corners and top middle of each flap are connected to a narrowed tissue using an absorbable suture. Here one may use the corner stitch mattress suturing technique (78). Next, the center and remaining intermediate points (Fig. 9G, middle panel) are connected together. The same is done for the opposite side.
At this point, PVA sponge implantation may take place. One PVA piece is inserted using Adson forceps underneath the P, M and D portions of each flap. Control sponges may be inserted in the outer area of a flap. Alternatively, a small pocket may be opened on the animal’s back.
A DPBS-moistened gauze is placed over the right flap. The left flap is positioned into place using a running suture technique with non-absorbable permanent suture (suggested directions of suturing are presented in Additional File 2, Fig. S7). The left flap is kept moistened and covered while the opposite side is sutured. Note: we do not recommend using proximate skin staplers with rotating head (e.g., Ethicon, #PRR35) to close the wound, especially in female rats that have thin and delicate skin.
The flaps and surrounding areas are wiped with moistened gauze to remove the traces of blood and betadine solution.
The animal is flipped into a prone position. 10 mL of saline is injected sc to supplement bodily fluids lost in the animal during this extensive surgical procedure.
Isoflurane flow is decreased to 0% so that the animal receives only 100% Oxygen.
The gross appearance of the flaps is recorded by digital photography (Fig. 9G, right panel).
Once the animal begins to awaken, we typically put a protective Elizabethan collar prior to returning the animal to a new padded cage with fresh food and water.
The HDPE tubes with collected tissue specimen are placed in -80⁰C freezer for storage.
All waste is removed, sharps disposed in specified containers, and all surgical instruments are washed, cleaned, and autoclaved at this point.

4.4 Post-surgical animal monitoring

When in distress, rodents can self-mutilate or cannibalize the ventral flap. To prevent self-destructing behavior, Elizabethan collars should be used instead of restraining jackets that cause immobility and thus will compromise flap healing. Although male rats adapt to these collars quicker than female rats, a careful pre-operative introduction of the collars to all animals is strongly advised. Despite the pre-operative preconditioning with the collar, female rats will often refuse to eat/drink after the surgery but will quickly pick up on any exposed sutures once the collar is removed. To avoid animal self-mutilation issues, many researchers choose to create the dorsal flaps as in the classical 1965 McFarlane experiment (79–83). However, one should note that dorsal flaps represent RPFs and thus a different flap healing model compared to the APFs.

Surgery-related stress can be reduced by introducing an animal to a habitat with novel enrichment devices and dietary treats. While animal socialization is otherwise certainly encouraged, the post-operative animals (and in general retired breeder males) should be housed separately so that the collars may be used safely, and animal food/water intake would be tracked more accurately. Food pellets, dough diet and/or nutritional special needs recovery diet gels (Bio-Serv) should be placed on the floor postoperatively for easy access. Rodent diet may also be medicated (Bio-Serv). Post-surgical animal well-being should be constantly monitored at least twice per day. Animal weight should be recorded every other day. Post-surgical pain relief is available if it does not interfere with the aims of the study as some pain-relieving therapeutic compounds are known to impede or accelerate an inflammatory response and change the gene/protein expression patterns. The suggested example of POAMS is provided as Additional File 1.

4.5 Non-survival surgery

4.5.1 Preparation for procedure

The HDPE tubes are labeled for appropriate specimen. A suggested label may contain a unique animal number, code for treatment, indication of left or right flap side (R/L), day of harvest and an abbreviated name of a harvested segment, e.g., “174871, A, L7D”).
10% buffered formalin container and histology cassettes are labeled for appropriate specimen.
A rubber bucket is filled with ice pellets. If PVA sponges were implanted, then 2.0 mL Eppendorf tubes are filled with PVA sponge lysis or other buffer, depending on the type of subsequent wound exudate (WE) analysis, and placed on ice. Flat-tip tweezers are prepared to handle the sponges.
Dewar flask is prefilled with LN₂ for immediate tissue freezing. We suggest using curved HDPE tube grabbing scissors.
CO₂ levels are checked in the euthanasia cart. Isoflurane and oxygen tank levels are checked in the anesthesia cart.

4.5.2 Flap harvest

The animal is placed in the anesthetic sedation chamber prefilled with 4–5% isoflurane gas and kept there for up to 4 minutes.
4% (larger rats) or 3% (smaller rats) isoflurane flow is redirected into the nose cone.
Once the animal is placed onto an underpad, which covers a heating pad, the digital photos of the flap appearance are taken. The functional outcome (full survival, partial failure, or total loss) of flaps exposed to SI should be evident by day 5, whereas flap fate following other treatments may be evaluated at later end-points. Figure 10A illustrates experimental situations, when the evaluation of flap survival percentage (FS%) by digital planimetry is not necessary (Fig. 10A, left panel – left flap, middle panel – both flaps) and when it is required (Fig. 10A, right panel – both flaps).
If the flaps are going to be divided into the proximal (P), medial (M) and distal (D) segments, a measuring ruler is placed on the top of a flap. Each flap is marked with either one dot, dotted or a solid line that would separate it into three equal portions. Note: this step should be performed on a living animal when the flaps are in fully extended position. If needed, the animal paws may be taped. If the IHC specimen is to be taken out of each portion, then two horizontal lines are drawn along the entire flap as shown in Fig. 10B.
All running sutures are disrupted and removed. One-by-one, the flaps are dissected along the wound margins and Mayo scissors are used to lift one flap after another from the wound bed. The status of the pedicle is inspected and POAMS form (Additional File 1) is used to indicate if it is functional or compromised. A fully functional vessel will be visibly dilated and pumping the blood (Fig. 10C). Obstructed vessel will have a dark blood clot or will be clear of the blood.
At this point, all PVA sponges may be retrieved if any were implanted previously. These sponges should be cleaned from the tissue prior to placing them into the pre-labeled Eppendorf tubes stored on ice. WE is collected though repeated compression of each sponge with flap-tip tweezers, and the samples are diluted if necessary.
If the flap is going to be divided into the P, M and D segments as shown in Fig. 10D, it should be preferably cut into the thirds while it is still attached to the pedicle. Alternatively, the entire flap may be cut at the base, transferred onto a sterile Petri dish kept on ice and divided over there. Optionally, the pedicle can be excised out of the P segment for a separate histological evaluation of other types of analysis. Whichever technique the surgeon chooses, he/she should stay consistent and adhere to the chosen sequence of flap harvest procedure throughout the entire course of experiment.
For IHC, the middle longitudinal strip is cut out of each flap’s segment (Fig. 10E). A specimen is placed into a dedicated IHC casette skin facing up. Edema will be the most prominent in the P segment of the flap, making it the thickest segment out of three. CRITICAL: the tissue strip should be laid straight and should not be twisted. The cassette is submerged in 10% formalin and incubated at RT for at least 48 h.
The remaining flap segment is placed in a pre-labeled HDPE tube, which is then submerged in LN₂. Even though the tissue may be small compared to the volume of the tube, it will less likely stick to the walls of the tube and will be easier to retrieve after freezing.
Other flap segments are processed in the same manner as described above.
For major vital organ harvest 5% (or greater) isoflurane flow is used to overdose the animal and then excise all organs of interest after its exsanguination. If organ harvesting is not required, then the gauze ios placed over both excisional wounds and the animal is moved into the clean cage for euthanasia. The CO₂ flow rate is adjusted at 5 L/min and exposure to CO₂ is continued for at least five minutes after respiratory arrest. Euthanasia may be confirmed by cervical dislocation (for rats < 200 g in weight) or bilateral thoracotomy. Note: other methods of euthanasia may be used as per approved IACUC study protocol.
Animal carcass is placed in a biohazardous waste bag, which is disposed of according to Institutional LAS rules. Typically, this entails attaching a label with a specific information, logging animal death, leaving a carcass in a dedicated freezer or cold room, and concluding a unique animal cage tag.
The surgeon’s working area is cleaned and sanitized. All surgical tools are washed, placed in the autoclave pouches/bags or trays, and then sterilized in the autoclave.

4.6 Data analysis

The quantitative and qualitative analytical techniques of the collected biological specimens include but are not limited to planimetric flap survival analysis, histology (HIS) (84), immunohistochemistry (IHC), laser capture microdissection (LCM) (85), flow cytometry (FC) (86), protein immunoblotting (IB) (87, 88), transcriptomic analysis through nucleic acid (NC), specifically mRNA, extraction, metabolomic analysis, and even isolation and analysis of large and/or small EVs from either tissues or wound fluids (89–91).

The principles of planimetric analysis are described in Additional File 2, Figs. S8 and S9. Total and either necrotic or viable areas in the digital photos can be determined with the aid of a “Measurement Log”, “Quick Selection” and “Ruler” tools available in Adobe Photoshop or ImageJ software. The percentage of flap viability or survival (FS%) is calculated as the necrotic flap area minus the total flap area divided by the total flap area (in square millimeters) multiplied by 100.

We also provide several simulated statistical flap survival data analysis examples in GraphPad Prism 9.5.0 software (Additional File 3 and Fig. S10). The differences between the matched left and right flap survival may be estimated by the paired Student’s T-test (Additional File 3, Example 1, and Fig. S10, upper left panel). The before and after treatment effects (e.g., baseline versus PI versus REP₁ vs SI) within each flap of different animals may be evaluated by one-way repeated measures (RM) ANOVA (Additional File 3, Example 2, and Fig. S10, upper right panel). For example, if the normalized phosphorylated Akt (p-Akt) protein expression effects were compared between the male and female animal groups consisting of 5 rats per each, we would be using two-way RM-ANOVA (Additional File 3, Example 3, and Fig. S10, bottom left panel). A good source of explanation when and how to use fully-nested ANOVA without interaction is provided by John H. McDonald in his topic “Nested Anova” (92), which implies that if we would have 7 randomly selected rats, with 2 flaps created within each under different ischemic conditions, and if we would carry out 4 independent skin oxygenation measurements (one before the flap raising surgery in the spots of future-to-be flaps and then on POD 2, POD 5 and POD 9) with three repetitive measurements per each segment of the flap each time, then our model could be analyzed either by “pure” Model II random-effects model-based nested ANOVA or by model I mixed-effects model-based nested ANOVA. In yet another example, if 14 male and 14 female non-littermate rats were randomly selected and assigned into 2 groups with 7 rats of each gender per each, and these groups would receive an external treatment (e g. tobacco vapors vs control water vapors) prior to BEFAF surgery, where one flap would be subjected to 4 h of PI alone and contralateral flap to 4 h PI, and then to 2 h of SI (e.g., AVO) after 12 h of REP₁, the nested ANOVA model would treat male and female rats as sub-groups and their flaps - as sub-sub-groups nested within these sub-groups. Furthermore, the distinct P, M and D segments of these flaps, if measured individually, would be treated as sub-sub-sub-groups. But if an overall flap viability % was measured at a single time point, e.g., POD 9, we could apply three-way 2x2x2 RM-ANOVA (Additional File 3, Example 4, and Fig. S10, bottom right panel). In the provided example, values stacked in sub-columns would represent a set of matched or repeated measures values (4 h PI vs 4 h SI). Means of cells that differ only by one factor would be compared using post-hoc Šidák test, reporting the multiplicity-adjusted P values for each comparison at alpha = 0.05 (95% CI). Residual, homoscedasticity and QQ plots would also be provided. An excellent feature of this type of analysis using GraphPad is the option to calculate cell/row/column means and grand means, which aid in plotting different results while considering or ignoring subject’s gender (see Additional File 3).

The tissue sections are quite large in size and there will be room to place only two replicate sections on each slide for HIS analysis. We prefer MT stain over H&E, as it readily reveals new and old collagen structures, extent of inflammatory infiltration, edema, and newly formed vessels. Depending on the extent of necrosis, the left and right sides of tissue section may appear heterogenous. If there is no possibility to digitize the slides using an automated or semi-automated hardware/software, then manual image collection via conventional or polarized light microscopy may be performed in a following manner: (a) the image of the entire flap segment is taken at the lowest (4x) magnification and medium to low sharpness; (b) the upper, middle, and lower sections of the P, M or D flap segments are taken at higher magnification (10x or 20x) making sure these sections slightly overlap; (c) The images are then combined together using Adobe Photoshop to produce one seamless high-quality image (Figs. 11 and 12).

Several images of the same area should be taken to reduce the risk of getting blurry poor-quality images (Additional File 2, Fig. S11). In our experience, the thickness of granulation tissue (GT) may be estimated from the 10x magnification micrographs (Fig. 13, left upper panel), while 20x and higher magnification micrographs are suitable for quantitative vessel analysis (Fig. 13, left bottom panel). While vascular density and angiogenic budding may be visualized in sectioned flap tissues stained with H&E (Fig. 13, left bottom panel) or MT dyes alone (Fig. 13, right upper panel), staining with specific endothelial phenotype markers such as PECAM-1, vWf and/or CD34 (Fig. 13, right bottom panel) may be preferred. Vascular inflammation that causes endothelial barrier disruption and tissue edema can also be studied using BEFAFs. Some proteins that are involved in endothelial barrier recovery, e.g. Hsp27 (93), are expressed in rat flaps (data not shown). 20x and higher magnification micrographs are suitable for the gross tissue cellularity analysis (Fig. 14, left panel), including the presence of mature adipocytes and tissue infiltration with nucleated hematopoietic cells, namely neutrophils and macrophages (identifiable by their characteristic polymorphic and euchromatic nuclei, respectively, and by their positive staining reaction with CD45) (Fig. 14, right panel). Collagen deposition and collagen fiber alignment can be examined at 10x magnification following MT or Sirius-Red staining (which can distinguish between different collagen types). Currently there are several automated free and open-source ImageJ and more sophisticated MATLAB software plugins created to analyze different types of scientific HIS/IHC images.

The isoleucine and hydroxyproline quantity in acid hydrolysates of each PVA implant will reflect total protein and collagen content. Specific cell populations can be isolated by FC from the WE filtered through a 70 µm cell strainer. For instance, a purified (> 95%) population of wound macrophages as determined by F4/80 staining can be yielded by magnetic cell sorting using mouse anti-CD11b tagged microbeads (94). A mixture of WE with specific lysis buffer to determine the latent and active matrix metalloproteinase (MMP) profile by gelatin zymography (95) and various growth factor/cytokine/chemokine levels by ELISA (96) or IB may provide critical information on the healing trajectory of a wound and its suitability for advanced biological therapies (97–100). Finally, one may choose to isolate proteins, mRNA or EVs from different flap segments using the materials and isolation procedures as suggested in Additional File 2.

Many tissue flap animal models have been developed and adapted to characterize disease pathophysiology, identify targets and evaluate the efficacy of novel therapeutic agents or treatment strategies (101). However, tissue flap models that aim to teach specific surgical skills and simultaneously study reparative processes still lack firm guidelines and standardization (102, 103). Our standardized reconstructive surgery model using BEFAFs shows promise as a more effective, less time consuming and cheaper alternative for validation of therapeutic targets, diagnostic and prognostic biomarkers as well as clinical treatment strategies than conventional wound healing animal models. This model is based on compound APF and can be readily modified to represent either pedicled or free flap under varied clinical situations. Unlike small excisional wounds, these flaps do not heal by contraction. Using microsurgical techniques, small caliber SIEA and SIEV vessels can be divided, and anastomosis can be performed under a microscope to directly simulate free flap surgery. Moreover, the procedure can be scaled and reproduced using other animal species (e.g., mice, rabbits, minipigs). One may assess the impact of endogenous or exogenous factors such as sex/gender, age, diet, physical activity, metabolic and chronic diseases on quantitative measures such as the rate of flap reoxygenation, vascularization, cell metabolism, migration, signaling and overall tissue survival. The BEFAF model is ideally suited to explore the molecular profiles associated with ischemic susceptibility or tolerance and to evaluate biomarkers for early prediction of flap outcome. Moreover, BEFAFs can be used to reveal effective therapies for prevention of flap loss or improvement of flap survival after salvage. For example, BEFAFs may be utilized to evaluate the biological effects of a preconditioning surgical delay technique, which triggers the expression and activation of angiogenic genes (104) and potentially stimulates cell-to-cell communication through the EVs. In this procedure, the left or right flap is raised, placed back onto the resultant wound, and left to fully heal for 7–14 days. Thereafter BEFAF surgery is performed on both sides of the animal creating any kind of flap. These flaps are allowed to heal under normal or perturbed conditions. Pre-surgical and post-surgical responses of interest as well as tissue survival are then compared between the control (acutely raised) and paired experimentally pre-conditioned (delayed) flap tissue at designated POD(s) (Fig. 15). Another example is application of exercise – the pre-surgical animals may be divided into two groups with sedentary and exercise regimen conditions, as described in our previous study using the rotational pedicled fasciocutaneous flaps (105).

Aside from an evaluation of flap healing rate, the animals may successfully undergo non-invasive in vivo microscopy. For instance, differences in left- and right-side flap vascularization and reoxygenation rates may be examined by combined photoacoustic and Doppler ultrasound imaging (Fig. 15), while dynamic metabolic changes in tissues may be assessed by intravital microscopy (83, 106). Temporal levels of intradermal nitrogen or oxygen may be measured by using specific needle-type microsensors or implants (107).

Finally, the BEFAF model can be used to evaluate a relative contribution of the recipient wound bed to APF or RPF survival. In such a case, a sterile silicone sheet is placed between the flap and the abdominal muscle. The sheet is sutured to the edges of a flap to prevent its movement, which may cause a physical damage to the pedicle (108). Although some wound bed isolation studies conducted on single animals showed a significant reduction of flap survival rates (109, 110), other investigators did not see a significant difference in functional flap outcome, but agreed that induction of potent pro-inflammatory response by a foreign material might conceal the true driving forces of the healing process (111). Ultimately, if superficial wound healing - not flap survival - is a preferable measure of outcome, then unipedicle or bipedicle ventral or dorsal bilateral flaps can be created to establish a gradual ischemic environment followed by punching a series of excisional wounds. These wounds may be further reinforced using silicone splinters to overcome the wound healing by contraction (112).

While the results of standardized studies using BEFAFs may be appropriate to guide recommendations for refining perioperative and post-operative flap management in specific human patient populations undergoing extensive reconstructive surgeries, this model presents challenges and has some limitations. In BEFAF, the survival/failure rate and efficacy of re-integration of right flap in response to the desired treatment regimen can be compared to that of saline-treated left flap control or vice versa. However, based on the results of our experiments using the methylene blue dye injection into the distal flap areas, the diffusion may impact the opposite side if the amount of injected solution exceeds 500 µL (in rats). It is also expected that the survival percent of left, and right untreated flaps may slightly vary. Although this naturally occurring within-subject variation typically does not exceed 10% of total flap area, an excessive manipulation of flap pedicles or flap corners can introduce unwanted heterogenous effects. Moreover, a degree of hypoxia induced in the distal portions of the flaps may differ not only between two different strains of animals but also between the same-strain animals that are received from different vendors. For instance, we noticed that Lewis strain rats have relatively thin blood with less time needed for reperfusion, thus their flaps had a better chance of survival than the same type flaps created in SD strain rats. In turn, SD strain rats obtained from Charles River Laboratories were more susceptible to flap failure after the SI compared to SD rats received from Envigo. Thus, for purely procedural repeatability studies, all surgeries should be preferably performed using littermate animals obtained from the same vendor.

We did not test whether BEFAF model can be applied to the clinical case of transposition flaps. Such flaps should be rotated towards the dorsal side of the animal, and it is unclear if their skin would be flexible enough to cover two simultaneously created excisional donor site wounds. Finally, in some experimental settings, rodents are exposed to repetitive anesthesia, which may induce respiratory failure and overdose-related death, especially in light weight animals. Susceptibility to overdose may also depend on animal gender: in our hands, female rats were less prone to overdosing than male rats. If the animal stops breathing during anesthesia, cardiopulmonary resuscitation is possible. Repetitive checking of animal vital signs, timely decrease, or cessation of anesthesia and sufficient time gaps between serial animal sedation events decreases fatal outcomes.

Accurate comparisons between standardized complex wound healing studies would add to the translational knowledge base of complex tissue healing under normal and pathological conditions. BEFAFs may be used to investigate the spatiotemporal cellular and molecular responses to complex tissue injury and interventions simulating clinically relevant flap complications (e.g., secondary arterial, venous, or mixed ischemia) and therapeutic or surgical treatments (e.g., flap delay) in the presence or absence of confounding risk factors (e.g., substance abuse, irradiation, diabetes) or favorable wound-healing promoting activities (e.g., exercise). This technically challenging but feasible reconstructive surgery model eliminates inter-subject variability, while concomitantly minimizing the number of animals needed to achieve adequate statistical power. Overall flap experiments designed for studying the effects of SI may be lengthy, require good time management skills, visual aids, awareness, and stamina. They may appear too laborious for novice trainees but have the merit of simulating real-life clinical surgeries sometimes lasting more than 8 hours. BEFAF also serves as an aid for teaching basic vascular microsurgery techniques that focus on technical and non-technical skill acquisition in the trainees.

7.1. Animals

Male and female 10–18-month-old retired-breeder Sprague-Dawley (SD) rats (Charles River Laboratories, Malvern, PA and Envigo, Indianapolis, IN) with an average weight of 310 g and 570 g were used in this study. Rats of same sex were acclimated at least 7 days prior to surgery under standard laboratory conditions (temperature 21°C ± 2°C, humidity 55% ± 10%, 12:12 h light-dark-cycle). Due to the weight (> 500 g) and retired breeder status males were housed singly. Females were housed in tandem. Food and water were provided in a standard manner. Some animals received liquid ethanol-supplemented diet.

7.2 General laboratory equipment and materials for the surgeon

This procedure used standard ACS or molecular biology grade (MBG) laboratory reagents/chemicals as well as standard veterinary surgery and microsurgery equipment/tools. In some instances, the shortened links to the randomly selected vendors that can supply specific surgical instruments or reagents used in this study are provided next to the item.

Personal protective equipment (PPE): surgical gown, ear loop face mask, hair cap, shoe covers, sterile surgical gloves, non-sterile gloves, safety glasses.
Stainless steel instrument tray with flap cover – for autoclaving and storage of surgical instruments
Stainless steel Grafco Flat Type instrument tray – for placing the surgical items during the procedure.
Sharps disposal container – for disposal of needles and syringes
Biohazard disposal container and biohazard bags – for safe disposal of biohazardous trash, including pipette tips.
Countdown timers
Digital camera with macro-view function

Optional:
Magnifying spectacles or loupes (2.5-3×, depending on the eyesight of the surgeon) with/without illumination or separate LED headlamp (> 500 lumens).

7.3 Materials for animal preparation for the surgery and flap design

Single or dual-channel anesthesia system with gas vaporizer, inlet flowmeter, charcoal filters, high pressure oxygen hose, oxygen regulator, and low-profile anesthesia masks adapted for rodents.
Small-animal scales
Secure animal transportation cart
Small animal heating pad
Handheld hair clipper and/or trimmer (e.g., WAHL for Home Pet). Note: The use of “Nair” or other chemically-based hair removal products is not recommended as they may interfere with the cutaneous gene/protein responses or cause allergic skin reactions.
Ophthalmic veterinary ointment (e.g., Puralube).
Betadine (10% povidone iodine) antiseptic solution.
Isoflurane inhalant anesthetic for veterinary use (USP)
Sterile non-woven or gauze sponges (e.g., Dermacea 4”×4”, 6-Ply by Covidien, #441405)
Disposable surgical under pads:

X-large 30”×36” Wings Ultra by Covidien.
Small 17”×24” or medium 23”×24” Dri-Sorb by Attends.

11. C-fold paper towels (e.g., Kleenex)

12. Sanitizing 70% ethanol spray

13. Sterile disposable skin marking pen with ruler (Viscot, #1422SRL9-100)
14. 1:3 or 1:2.5 ratio flap design template made from sturdy card-paper with extra 1–2 cm in length if distal biopsy collection is intended.

7.4. Materials for major or minor survival surgery

Pre-operative analgesic (opioids such as sustained release (SR) buprenorphine HCl are typically obtained from LAS veterinarian staff). Otherwise, take a new sterile 1 ml syringe, slowly draw in analgetic solution into the syringe. Point the syringe upward to bring any air bubbles to the top of the needle. Plunge the syringe slowly to expel any air until only liquid expels from the syringe. Double check volume, place the needle cap and set a syringe aside.
Sterile Dulbecco's Phosphate-Buffered Saline (DPBS) (Cytiva, #SH30028.02) or HyClone™ Hank's Balanced Salt Solution (HBSS) without calcium, magnesium, and phenol red (Cytiva, #SH30588.01) – for irrigation of raised flaps while maintaining cell tonicity and viability.
1% Lidocaine HCl - for topical irrigation of pedicle to prevent or reduce traumatic vasospasm(s). Prepare 1% solution using sterile DPBS.
10 mL saline pre-filled syringes (e.g., BD PosiFlush™ by BD Biosciences).
1 mL sterile disposable syringes with Luer-Lok tips
Sterile 21 gauze hypodermic needles
Hot glass bead dry sterilizer (e.g., Germinator 500 by BrainTree Scientific) – for surgical instrument sterilization in between two successive animals that undergo flap raising surgery on the same day.
Multi-Enzymatic Cleanser – for surgical instrument cleaning (McKesson, #484478)
Sterile surgical instruments (available from Electron Microscopy Sciences (EMS; Hatfield, PA), Kent Scientific Corporation (KSC; Torrington, CT), Fine Science Tools (FST, Foster City, CA), World Precision Instruments (WPI; Sarasota, FL) or other commercial sources):

Iris Supercut scissors (curved or straight) - for tissue cutting.

Example

https://tinyurl.com/5n7r8mez

Metzenbaum-Nelson scissors (straight) – for excisional biopsy cutting.

Example

https://tinyurl.com/3jmc5r8d

Mayo dissecting scissors (curved with beveled blades, medium or large size) – for tissue dissection.

Example

https://tinyurl.com/3p3nnb68

McPherson Vannas or Noyes Iris micro scissors (straight with beveled smooth blunt/blunt or sharp/sharp blades) – for dissection of tissue around pedicle and vessel separation.

Example

https://tinyurl.com/2p8mfwc9

Halsey micro needle driver/holder with serrated tips – for holding a needle during vessel separation.

Example

https://tinyurl.com/mvm7nepx

Kelly Hemostat forceps (straight with serrated tips) – for suture needle grasping and holding.

Example

https://tinyurl.com/3trzus3n

Adson tissue forceps with 1x2 teeth – for lifting tissue during suturing.

Example

https://tinyurl.com/6kbsap4m

Micro/fine forceps (slightly and fully curved, smooth jaws) – for lifting the pedicle and fascia.

Example

https://tinyurl.com/yckfr564

Harms micro suturing (straight) and Mcpherson tying forceps (angled) – for performing anastomosis (optional)

Example

https://tinyurl.com/422r2252 and https://tinyurl.com/bddtpus3

Flat-tip tweezers with upward bent tips (EMS #78336-36A) – for handling PVA sponges

Example

https://tinyurl.com/33rbmyfv

10. Atraumatic vascular clamps:

7 mm B-1V, 3.5 x 1mm (FST, #00396-01) – for vein occlusion
11 mm B-2V, 5.5 x 1.5mm ((FST, #00398-02) – for artery occlusion and global ischemia
16 mm – B-3V, 7.5 x 1.75mm (FST, #00400-03) for occlusion of larger diameter vessels in larger laboratory animals.

Example

https://tinyurl.com/yc22wa4m

11. Clamp applying forceps with lock (FST, #00071 − 14) – for handling vascular clamps.

Example

https://tinyurl.com/2bvx2zpf

12. Disposable or reusable battery-operated high temperature cautery with fine tip (e.g., Bovie Medical #AA01; EMS #72976-31) – to prevent bleeding during microvascular tissue dissection.

13. Sterile 4 − 0 size sutures with 16 mm (for female rats) or 19 mm (for male rats) 3/8 circular reverse cutting needle:

Non-absorbable 18” or 30” POLYPROPYLENE or NYLON – to place running sutures.
Non-absorbable 18” or 30" SILK – to place temporary sutures and vascular separation.
Absorbable 18” or 30" VICRYL Polyglactin 910 – to place buried simple-interrupted sutures.

14. Elizabethan collars (small, medium, large, or x-large)

15. Small or medium size LIGACLIP® Multiple-Clip Applier (MCA) – for permanent occlusion of vessels (e.g., Ethicon, #MCS20 or #MCM20)

Optional:

16. 10 mL BD PosiFlush™ Pre-Filled Heparin Lock Flush Syringes (100U/mL) – for topical dropwise irrigation of vascular lumen prior to vascular occlusion (BD Biosciences, #306512)

7.5 Materials for tissue biopsy and flap excision

Liquid nitrogen (LN₂) supply and portable LN₂ dewar (e.g., 4L by Nalgene)
Cryogenic flat round-tip tweezers
18 mL polyethylene scintillation HDPE vials with threaded cap (e.g., Fisher Scientific, # 14-955-393) – for tissue sample storage and easy access
Fine-tip histology marker and LN₂-proof cryo-marker (e.g., Dual-point UltraCruz, Santa Cruz Biotechnologies,# sc-360977)
-80⁰C freezer – for long-term tissue and sample storage.
Animal euthanasia cart with CO₂ gas tank supply
Small biohazardous waste bags or specified animal carcass disposal bags

Optional: Sterile Disposable McKesson Biopsy Punches (available in 2–10 mm sizes)

7.6 Tissue preparation for histology

Unisette™ 1-chamber histology embedding/processing cassettes with 1 mm openings (Ted Pella) - for smaller size and leaner specimens:

Outside dimensions: 28.5 x 41 x 6.7mm (1.12" x 1.61" x 0.26").
Inside dimensions: 26 x 30 x 5mm (1.02" x 1.18" x 0.2").

Macrosette™ 1-chamber histology embedding/processing cassettes with 1 mm openings (Ted Pella) - for thicker specimens
Micromesh™ 4-compartment biopsy cassettes – for excised pedicles
Cassette marking histology pen (e.g., Statmark by StatLab)
90 ml containers with prefilled 10% Neutral Phosphate-Buffered Formalin (pH 7.2 ± 0.5; vials) (e.g., StatLab, # NB0345)
1X PBS buffer without magnesium and calcium - for cassette washing after fixation. Store at RT.
70% Ethanol solution – for cassette storage until analysis at + 4⁰C.
Large glass staining dishes.
Stains of interest, e.g., Masson’s Trichrome (MT) and Hematoxylin & Eosin (H&E).
Classical histology lab equipment if not using Core Histology facility

7.7 Other Materials

Materials for an optional isolation of tissue-derived proteins, mRNA, extracellular vesicles (EVs), and materials for optional PVA sponge implantation and processing to extract wound exudate (WE) material are provided in the Additional File 2.

11.1 Ethics approval

All animal experiments were conducted in accordance with ARRIVE guidelines as well as policies of the NIH Guide for the Care and Use of Laboratory Animals (NIH Publications No. 8023, revised 1978) and were approved by the Institutional Animal Care and Use Committee (IACUC) of Thomas Jefferson University (TJU) (IACUC protocols #01713, #01481, #020181).

11. 2 Availability of data and materials

Data sharing is not applicable to this article as no datasets were generated or analyzed during the current study.

11. 3 Competing interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

11. 4 Funding

This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors, and was supported by the Department of Otolaryngology – Head & Neck Surgery, Thomas Jefferson University, Philadelphia, PA.

11. 5 Authors' contributions

EP: Conceptualization, Methodology, Project Administration, Resources, Funding acquisition, Writing - Reviewing and Editing. RH: Methodology, Project Administration, Supervision, Writing - Reviewing and Editing. JH: Supervision, Resources, Writing - Reviewing and Editing. EA: Investigation, Visualization, Formal analysis, Writing - Original draft preparation.

11. 6 Acknowledgments

The authors are grateful for excellent technical support, guidance and animal care received from TJU Department of Laboratory Animal Services staff, and especially thank Veterinary Technician Evelyn Skoumbourdis, MS, RLATG as well as animal caretaker Victoria Jackson. The authors thank Dr. John R. Eisenbrey and sonographer Maria Stanczak, MS, RDMS, RVT for their services providing animal PA-DUS imaging, and TJU’s Translational Research/Pathology Core facility manager Dr. Zhijiu Zhong and his staff for their excellent histological stain services.

11. 7 Authors' information (in alphabetical order)

EA is a long-term member of Wound Healing Society (WHS), American Society for Biochemistry and Molecular Biology (ASBMB) and American Society for Investigative Pathology (ASIP). She completed her undergraduate and postgraduate studies in the Faculty of Natural Sciences at Vytautas Magnus University (VDU) (Kaunas, Lithuania), and received the B.S. in Biology (2001), M.S in Molecular Biology & Biotechnology (2003) and PhD in Biochemistry (2007) degrees from VDU’s Department of Biology. She joined Prof. Boris N. Kholodenko’s Systems Biology and Cell Signalling Networks research group at the Department of Pathology, Anatomy and Cell Biology at Thomas Jefferson University (TJU) (Philadelphia, PA, USA) as a Research Assistant in 2004 to study the receptor tyrosine kinases (RTK)-mediated cell signalling. While at TJU, she completed her postdoctoral research in Cancer Signal Transduction as a NIH NIAAA T32 grant-funded trainee. In 2014, EA joined the Department of Otolaryngology – Head and Neck Surgery (TJU) to serve as a Research Instructor. She continued her research investigating cellular pathway interactions in reparative wound healing, and mentored Department’s residents teaching them basic biochemical analysis methods and surgical animal model techniques. Currently, EA holds a Research Scientist position at the Beckman Institute for Advanced Science and Technology, which is a unit of University of Illinois Urbana-Champaign (Urbana, IL, USA) dedicated to interdisciplinary research. She coordinates human clinical research studies in Prof. Stephen A Boppart’s Biophotonics Imaging Laboratory that collaborates with a pharmaceutical industry company GSK (formerly GlaxoSmithKline) and takes part in projects focused on human ear diseases and cancer. She has recently joined the International Society for Extracellular Vesicles (ISEV) and is actively studying cancer-derived extracellular vesicles and their signalling roles.

EDP is a Professor at Department of Otolaryngology – Head and Neck Surgery at Thomas Jefferson University (TJU) (Philadelphia, PA, USA), an Executive Vice President at Jefferson Health and the President of the Jefferson Medical Group. He graduated from Perelman School of Medicine at the University of Pennsylvania in 1986 and completed his residency in Hospital of University of Pennsylvania (HUP). He received a fellowship from Stanford University Hospital (Stanford, CA) and is Board-Certified in Otolaryngology as well as Facial Plastic and Reconstructive Surgery. EDP holds active ENT physician appointments at Thomas Jefferson University Hospital (TJUH) and Methodist Hospital Division of TJUH (Philadelphia, PA). He is interested in selectivity and response characteristics of human olfactory neurons which would help identifying treatment for taste and smell disorders. He also specializes in thyroid carcinoma and thyroid, parathyroid and rhinoplasty surgery. EDP has been a consecutive P-50 NIH Grant Co-Investigator, served on the Sidney Kimmel Medical College Dean's Diversity, Equity & Inclusion (DEI) Council (Philadelphia, PA) and led the Educational and Research Foundation for the American Academy of Facial Plastic and Reconstructive Surgery (AAFPRS) Domestic Violence Project. EDP serves on the Board of Trustees of Wilmington University (New Castle, DE, USA). He and Patrizio Cernetti co-founded medical devices company, which has patented a drug delivery device that attaches to any smart phone. He wrote the first BoTox dose response paper and originated the Sonic Rhinoplasty technique. As a leader and mentor adept at building strong cross-functional teams, EDP continuously supports medical student scientific research rotation programs teaching how to translate various pre-clinical surgical animal models to human clinical research.

JBH is a Professor and Vice-Chair for Research at Department of Pathology, Anatomy and Cell Biology at Thomas Jefferson University (TJU) (Philadelphia, PA, USA). JBH trained in biochemistry with Drs. J.M. Tager and E.C. Slater at the University of Amsterdam, focusing on mitochondrial metabolism. He received his Ph.D. degree in 1972. During this time, he also worked with Dr. H.A. Krebs in Oxford, UK, on the integration of mitochondrial and tissue redox metabolism. He then pursued postdoctoral studies with Dr. Lars Ernster in Stockholm, Sweden, after which he joined the faculty of the University of Nairobi in Kenya. Dr. Hoek was introduced to alcohol research in 1981, when he joined the group of Dr. Emanuel Rubin at Hahnemann University in Philadelphia. The alcohol research group moved in 1986 to TJU. JBH has held numerous leadership and advisory positions in the alcohol research field. He has served on several NIH and NIAAA study sections, and from 2004 to 2008 was a member of the NIAAA National Advisory Council. He is currently a member of the Medical Advisory Council of the Alcoholic Beverages Medical Research Foundation (ABMRF). JBH is Associate Editor for Reviews and Commentaries for Alcoholism, Clinical and Experimental Research. He also chairs the Publication Committee of the Research Society on Alcoholism. JBH is engaged in multiple scientific collaborations in the U.S. and internationally. He serves on the steering committee of the German Liver Systems Biology consortium "HepatoSys," and he works closely with the newly established Systems Biology Center at University College Dublin, Ireland. NIAAA has honoured JBH with the Mark Keller Award. The research in interdisciplinary JBH’s laboratory focuses on the unique regenerative capabilities of the liver and how they are distinctively regulated by mitochondrial calcium signalling and alcohol metabolism. JBH’s laboratory developed different rodent alcohol consumption models to analyse the integrated signalling responses in liver and other tissues and study the adaptive or maladaptive modification of these responses by acute and chronic ethanol exposure. Some of his models were adapted to investigate the impact of alcohol on ischemic wound healing.

RNH is a Professor at Department of Otolaryngology – Head and Neck Surgery at Thomas Jefferson University (TJU) (Philadelphia, PA, USA. He received his undergraduate degree from Yale University (New Haven, CT, USA), and began his medical education at Jefferson Medical College (Philadelphia, PA) where he received his MD degree. He was elected to the Alpha Omega Alpha Medical Honour Society and was awarded the S. MacCuen Smith Memorial Prize in Otology. RNH completed his internship in General Surgery followed by a surgical residency and specialization in Otolaryngology – Head and Neck Surgery at TJU Hospital (Philadelphia, PA). RNH was then chosen for a highly specialized Fellowship in Facial Plastic and Reconstructive Surgery, sponsored by the American Academy of Facial Plastic and Reconstructive Surgery (AAFPRS) at the University of California (Los Angeles, CA, USA). He returned to TJU in 2006 and became the Director of the Division of Facial Plastic and Reconstructive Surgery. He is highly active in teaching the latest advancements and techniques in plastic surgery to residents and fellows. RNH is Double Board-Certified Facial Plastic and Reconstructive Surgeon at TJU Hospital (Philadelphia, PA) and a fellow of the American Academy of Surgeons. As a facial plastic surgeon, RNH specializes solely in plastic surgery of the face, head, and neck, including facial nerve paralysis treatment, corrective surgery for facial defects, microsurgery, reconstruction of cancer patients and extensive post-traumatic deformities. RNH has been named a Castle Connolly Top Doctor and a Philly Top Doc yearly since 2017. He is one of five surgeons regularly serving the humanitarian Children’s Rehabilitation Institute and Surgery Program (C.R.I.S.P.).

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No competing interests reported.

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Editorial decision: Major revision
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Novel, standardized pre-clinical surgical animal model to investigate the cellular and molecular mechanisms of ischemic flap healing

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Journal Publication

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Abstract

Figures

BACKGROUND

RESULTS

4.1. Overview of BEFAF model

4.2 Experimental animal preparation for survival surgery

4.3 Major survival surgery

4.3.1 Preparation

4.3.2 Flap design

4.3.3 Flap raising

4.3.4 Physical vessel separation and selective clamping to induce primary ischemia (PI)

4.3.5 Partial flap closure

4.3.6 Primary reperfusion (REP1)

4.3.7 Secondary ischemia (SI)

4.3.8 Secondary reperfusion (REP2)

4.3.9 Full flap closure

4.4 Post-surgical animal monitoring

4.5 Non-survival surgery

4.5.1 Preparation for procedure

4.5.2 Flap harvest

4.6 Data analysis

DISCUSSION

CONCLUSIONS

MATERIALS AND METHODS

7.1. Animals

7.2 General laboratory equipment and materials for the surgeon

7.3 Materials for animal preparation for the surgery and flap design

7.4. Materials for major or minor survival surgery

7.5 Materials for tissue biopsy and flap excision

7.7 Other Materials

DECLARATIONS

REFERENCES

Additional Declarations

Supplementary Files

Status:

Journal Publication

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4.3.6 Primary reperfusion (REP₁)

4.3.8 Secondary reperfusion (REP₂)