1. Art and Diseño

ORIGINAL ARTICLE
In Vivo Evaluation of Wound Bed Reaction and Graft
Performance After Cold Skin Graft Storage: New
Targets for Skin Tissue Engineering
Alicia Knapik, MSc,* Kai Kornmann,* Katrin Kerl, MD,† Maurizio Calcagni, MD,*
Christian A. Schmidt, MD, PhD,‡ Brigitte Vollmar, MD,§ Pietro Giovanoli, MD,*
Nicole Lindenblatt, MD*
Surplus harvested skin grafts are routinely stored at 4 to 6°C in saline for several days in
plastic surgery. The purpose of this study was to evaluate the influence of storage on human
skin graft performance in an in vivo intravital microscopic setting after transplantation.
Freshly harvested human full-thickness skin grafts and split-thickness skin grafts (STSGs)
after storage of 0, 3, or 7 days in moist saline at 4 to 6°C were transplanted into the
modified dorsal skinfold chamber, and intravital microscopy was performed to evaluate
vessel morphology and angiogenic change of the wound bed. The chamber tissue was
harvested 10 days after transplantation for evaluation of tissue integrity and inflammation
(hematoxylin and eosin) as well as for immunohistochemistry (human CD31, murine
CD31, Ki67, Tdt-mediated dUTP-biotin nick-end labelling). Intravital microscopy results
showed no differences in the host angiogenic response between fresh and preserved grafts.
However, STSGs and full-thickness skin grafts exhibited a trend toward different timing
and strength in capillary widening and capillary bud formation. Preservation had no
influence on graft quality before transplantation, but fresh STSGs showed better quality
10 days after transplantation than 7-day preserved grafts. Proliferation and apoptosis
as well as host capillary in-growth and graft capillary degeneration were equal in all
groups. These results indicate that cells may activate protective mechanisms under cold
conditions, allowing them to maintain function and morphology. However, rewarming may
disclose underlying tissue damage. These findings could be translated to a new approach
for the design of full-thickness skin substitutes. (J Burn Care Res 2013;XXX:00–00)
Reconstructive surgery applies several methods to
cover acute or chronic wounds. For skin defects,
current options for coverage include not only the
From the *Division of Plastic and Reconstructive Surgery, Department of Surgery, University Hospital Zurich, Switzerland;
†Division of Dermatology, University Hospital Zurich, Switzerland; ‡Clinic for Cardiovascular Surgery, University Hospital
Zurich, Switzerland; and §Institute for Experimental Surgery,
University of Rostock, Germany.
This study was financially supported by the Swiss National Science
Foundation (SNF-Grant 310030_127366). The authors declare
that they have no competing financial interests, commercial
associations, or financial disclosures that might pose or create a
conflict of interest with information presented in the article. The
authors disclose funding from NIH, Welcome Trust, HHMI.
Address correspondence to Nicole Lindenblatt, MD, Division of
Plastic and Reconstructive Surgery, Department of Surgery,
University Hospital Zurich, Raemistrasse 100, 8091 Zurich,
Switzerland.
Copyright © 2013 by the American Burn Association
1559-047X/2013
DOI: 10.1097/BCR.0b013e3182a226df
standard procedure, which is split- or full-thickness
skin grafting,1 but also the use of skin substitutes.2
Products currently applied and commercially available in clinical routine are acellular collagen dermal
matrices (eg, Integra®, Matriderm®) or epidermal
substitutes (Epicel®, Epibase®). Skin substitutes were
extensively reviewed by Shevchenko et al3 and most
of them specifically lack properties of full-thickness
skin, that is, color match, skin appendices, pliability, elasticity, and most of all mechanical stability.4 It
is obvious, that until today no successful full-thickness skin substitute design has been achieved. The
presently available full-thickness constructs usually
acquire only insufficient blood supply, leading to
cell death in the substitute. Strategies to overcome
this problem include prevascularization of skin substitutes,5 bioprinting and electrospinning of biomaterials including application of defined growth
1
Copyright © American Burn Association. Unauthorized reproduction of this article is prohibited.
Journal of Burn Care & Research
Month/XXX 2013
2 Knapik et al
factor cocktails,6 and finally the use of stem cells.7,8
All these strategies aim at speeding up the time until
skin substitutes obtain sufficient blood supply and
cellular composition. In the clinical setting, surplus
harvested skin grafts are commonly stored at low
temperatures (4–6°C) in order to make use of them
if rest defects occur.9 However, nowadays clinical
institutions have to follow rules with regard to good
clinical practice and good manufacturing practice.
The European Parliament issued the 2004/23/EG
guidelines in 2004 in order to improve tissue quality and to increase patient safety in the setting of
human tissue preservation. These guidelines define
the quality and safety standards required for donation, procurement, testing and processing, and preservation, storage, and distribution of human tissues
and cells. The guidelines have a major impact on all
aspects of tissue banking and transplantation and
demand standardized tissue storage under controlled
conditions.10,11
In a previous study we evaluated the effect of cold
storage in moist gauze at 4 to 6°C on split-thickness
skin graft (STSG) tissue integrity and viability, cell
proliferation, apoptosis, and vascularization.12 Histologic findings showed a preserved integrity of the
tissue over a period of 7 days. However, metabolic
activity of the tissue decreased by 50% after day 3
of storage, indicating a slowed down oxygen consumption and viability. Metabolic activity stayed at
this level until 14 days of storage at 4 to 6°C. Thus,
stored skin grafts adjusted their metabolic demand
under cold temperatures, prolonging survival without blood perfusion. The concept of lowering nutritional demand by lowering temperature is a known
phenomenon in nature, namely it is encountered
during hibernation in mammals.13 Many of the physiological extremes of hibernation would be expected
to lead to apoptosis or necrosis in nonhibernating
animals.14 The knowledge whether similar processes
can potentially take place on human tissue—in this
case in skin grafts—may open interesting possibilities
to induce these mechanisms in skin substitutes. By
this, the survival of skin substitutes grafted onto the
human body could be prolonged during rewarming from the chilled and metabolically reduced state
before complete acquisition of blood supply has
been achieved.
Therefore, it is the aim of this study to investigate the effect of storage of human STSGs at 4
to 6°C up to 7 days on their in vivo performance
after transplantation in the modified dorsal skinfold
chamber in SCID mice.15 Angiogenic reactions of
the wound bed, capillary regression, and ingrowth
of wound bed vessels into the graft were accounted
for. Moreover, graft performance was determined by
histological assessment.
METHODS
Patients and Split-Thickness Skin Graft/FullThickness Skin Graft Harvest
The study was approved by the local ethics committee. Written and informed consent was given by
every skin donor (n = 19). The age of the patients
ranged from 28 to 71 years. Patients were excluded
in case of skin illnesses. Healthy patients underwent
different procedures of dermolipectomy (abdominoplasty, belt lipectomy, and thigh lift). STSGs of
300 µm thickness were taken from the specimen
with an air-driven dermatome (Zimmer, Münsingen, Switzerland) and directly transplanted or
stored for 3 and 7 days in saline-moisturized gauze
at 4 to 6°C in the refrigerator. Full-thickness skin
grafts (FTSGs) were harvested with a scalpel separating the dermis from the underlying fat layer and
transplanted directly.
Mouse Modified Dorsal Skinfold Chamber
On approval by the local government, all experiments were carried out in accordance with the Swiss
legislation on protection of animals and the EC
directive 86/609/EEC for animal experiments. All
animals received humane treatment, were housed in
separate cages with a 12-hour light/dark cycle, and
given water and food ad libitum. Microcirculation
was studied in the modified dorsal skinfold chamber as described previously.15 B6.CB17-Prkdcscid/
SzJ mice (“SCID” mice, n = 18; Jackson Laboratories, Bar Harbour, ME) with a body weight of 23 to
27 g were anesthetized by an intraperitoneal injection of ketamine (90 mg/kg body weight) and xylazine (25 mg/kg body weight). Briefly, for chamber
implantation, two symmetrical titanium frames were
mounted on a dorsal skinfold of the animal. One
skin layer was then completely removed in a circular
area of 15 mm in diameter, and the remaining layers (consisting of striated skin muscle, subcutaneous
tissue, and skin) were covered with a glass coverslip
incorporated into one of the titanium frames. Before
skin grafting, the animals were allowed a recovery
period of 3 days. Then, skin and most parts of the
hypodermal fat layer were carefully removed in a circular area of 7 mm in diameter from the back of the
chamber, leaving the panniculus carnosus as wound
bed. Because of technical reasons it was not possible to harvest STSGs in a reproducible way, and
with comparable thickness from the highly elastic
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Journal of Burn Care & Research
Volume XXX, Number XXX
Knapik et al 3
Figure 1. A, Intravital microscopy of the wound bed. Schematic view of the respective areas studied in the center and the
periphery of the wound bed. B, Areas of histological determination of proliferation and apoptosis rate in the upper dermis
(1–3), lower dermis (4–6) of the graft, and the host wound bed (7–9).
and thin mouse skin. Therefore, we chose to transplant human skin grafts. Three different groups were
defined (n = 5 per group): 1) freshly harvested human
STSGs; 2) 3-day preserved STSGs; and 3) 7-day preserved STSGs. Further SCID mice (n = 3) received
directly transplanted human FTSGs to compare the
wound bed reaction with STSGs. Grafts were placed
into the defect in the back of the chamber. By this, it
was possible to visualize the microcirculation of the
wound bed in vivo and to study the morphological
changes of the vasculature resulting as a response to
graft placement.
Intravital Microscopy
Repetitive intravital microscopic analyses of wound
bed and skin graft were carried out regularly during a period of 10 days. Because of the density and
thickness of human epidermis including a thick
layer of keratinocytes, it was not possible to monitor the microvasculature in the graft itself. Microscopic images were taken at eight different areas
within the center and the periphery of the wound
bed (Figure 1A). After injection of 0.15 ml fluorescein isothiocyanate–labelled dextran (1%; MW
70000; Sigma-Aldrich, Munich, Germany) the
microcirculation was visualized by intravital fluorescence microscopy (AxioScope.A1; Zeiss, Feldbach, Switzerland). Microscopic images were
captured by a CCD camera (AxioCam HSm; Zeiss)
using ×10 (N-Achroplan 10x/0,25, Zeiss) and ×20
(W N-Achroplan 20x/0,5; Zeiss) objectives in
AxioVision Rel 4.8. Vessel morphology and angiogenic changes were monitored in capillaries of the
muscular wound bed.
Microcirculatory Analysis
Microvascular perfusion was quantified offline by
analysis of the digital images using a computerassisted image analysis system (CapImage; Zeintl
Software, Heidelberg, Germany). This included the
characterization of morphological vascular changes
(vessel diameter, capillary density, and formation of
angiogenic buds).
Histopathological Evaluation of Tissue
Integrity
Ten days after transplantation the chamber content
was harvested in total (ie, skin graft attached to
wound bed) and fixed in 4% formalin and subsequently embedded in paraffin, according to standard procedures. Sections (4 µm) were stained with
hematoxylin and eosin staining for general morphological aspect. Samples were scored concerning
microscopic appearance by a dermatopathologist in
a blinded fashion. The scoring system was adapted
from the study by Başaran et al (Table 1).9 The
score estimates morphologic changes of the epidermis and dermis as well as collagen orientation,
inflammation based on leukocyte infiltration, and
apoptosis of keratinocytes. The total score was calculated by adding the appropriate separate score of
each parameter, resulting in a possible maximum
score of 10.
Table 1. Parameters for histological assessment of graft
quality
Microscopic
Parameter
Epidermal integrity
Epidermal–dermal
junction
Collagen
organization
Inflammation
Apoptotic
keratinocytes
Score
0
1
2
Destroyed
Destroyed
Partial
Partial
Normal
Normal
Amorph
Disturbed
Normal
Complete
Complete
Similar to control
Present
None
Normal
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Journal of Burn Care & Research
Month/XXX 2013
4 Knapik et al
Immunohistochemical Evaluation of Human
and Mouse Capillary Density
Paraffin sections (4 µm) were stained with mouse
antihuman CD31 (Dako, Glostrup, Denmark) or
rat antimouse CD31 (Dianova, Hamburg, Germany), incubated and detected with ABC-Vectastain (Reactolab, Servion, Switzerland) stained
using AP substrate or DAB chromogen (Dako),
respectively. The antigen retrieval for human CD31
staining required a Proteinase K (Dako) digest,
whereas the preparation for retrieval of mouse
CD31 required a treatment in boiling ethylenediaminetetraacetic acid buffer (pH 9). Slides were
scanned with the Mirax Midi Slide Scanner (Zeiss)
under a ×20 dry objective. Pictures were taken with
a 3CCD colour camera (1360 × 1024 pixels). Capillaries were counted in five random dermal areas of
cross-sections of the skin.
Immunohistochemical Assessment of Cell
Proliferation
Paraffin embedded sections (4 µm) were singlelabelled with rabbit antihuman Ki67 (Abcam, Cambridge, United Kingdom) after antigen retrieval in
boiling citrate buffer, followed by antibody incubation and detection using ABC-Vectastain (Reactolab). Quantification of stained cells was performed
by counting nine representative fields for each skin
section in the upper dermis, deeper dermis, and
the muscle using a ×40 objective (Figure 1B). The
proliferation rate is expressed as the ratio of the
number of proliferating cells divided by the total
cell number.
Determination of Cell Apoptosis Rate
Apoptosis was assessed by in situ Tdt-mediated
dUTP-biotin nick-end labelling, using a commercially available kit (Qbiogene, Illkirch, France).
Quantification of stained cells was performed by
counting representative fields for each skin in the
lower dermis, deeper dermis, and the muscle section
using a ×20 objective (according to Figure 1B); the
apoptotic cell rate is given as proportion of the number of positive cells per total number of cells.
Statistical Analysis
Differences between values were assessed using oneway analysis of variance or Kruskall–Wallis tests followed by the appropriate post hoc comparison tests
(Holm–Sidak and Dunn’s test, respectively). All data
were expressed as means ± SEM and overall statistical significance was set at P <.05.
RESULTS
Comparison Wound Bed Reaction After
Human Split-Thickness Skin Graft and FullThickness Skin Graft Transplantation In Vivo
In order to exclude differences resulting from the
nature of the placed graft, experiments comparing
the angiogenic host response after FTSG and STSG
placement were performed. Overall, there were
no significant differences in wound bed reactions
between STSG and FTSG. However, there was a
tendency toward a different timing of host angiogenic response and its magnitude (Figure 2 A–C).
After FTSG placement, host capillaries replied within
3 days with an extensive capillary widening, whereas
the placement of an STSG led to a weaker and
delayed response starting after 5 days. Host capillaries showed an early (day 3 posttransplantation) and
strong bud formation in response to STSG, but a
weaker and later (day 5 posttransplantation) reaction
after FTSG placement. Capillary density was slightly
higher after STSG placement and showed a similar
development over time when compared with that
after FTSG placement.
Influence of Split-Thickness Skin Graft
Preservation Time on Wound Bed Response
In Vivo
Intravital microscopy revealed that the host angiogenic response did not show any differences after the
different storage periods. Neither capillary density
(Figure 3A), nor capillary diameter (Figure 3B) or
bud formation (Figure 3C) were significantly influenced by the time of graft storage. Capillary density
for all three storage periods was in the normal range
of the dorsal skinfold chamber (250–350 cm/cm2)
and dropped slightly after transplantation. Angiogenic response symbolized by bud formation started
after day 2 and regressed until day 10 in a similar
manner in the three experimental groups. Capillary
widening was weak and occurred after 6 days, comparable in all three groups.
Influence of Preservation Time on Graft
Integrity
Skin grafts preserved for 7 days and transplanted into
the dorsal skinfold chamber exhibited a significantly
lower total graft score after 10 days than freshly
transplanted skin grafts (Figure 4A and B). However, no significant difference in the graft score after
storage of 0, 3, and 7 days was observed after 10
days, indicating that graft storage up to 7 days does
not affect mere tissue integrity after transplantation.
Copyright © American Burn Association. Unauthorized reproduction of this article is prohibited.
Journal of Burn Care & Research
Volume XXX, Number XXX
Knapik et al 5
C
Capillary density (cm/cm2)
400.00
Capillary diameter (um)
Human FTSG
300.00
250.00
200.00
150.00
100.00
50.00
0.00
B
Human STSG
350.00
0
3
5
7
10
Angiogenic wound bed reaction (buds
n/mm2)
A
16.00
Human STSG
Human FTSG
14.00
12.00
10.00
8.00
6.00
4.00
2.00
0.00
0
14
3
5
7
10
14
Days after Tx
Days
25.00
Human STSG
Human FTSG
20.00
15.00
10.00
5.00
0.00
0
3
5
7
10
14
Days after Tx
Figure 2. Comparison of the wound bed reaction between STSGs and FTSGs transplanted into SCID mice. Capillary density (A) and capillary diameter (B) in wound bed. Angiogenic response of the host (C). Means ± SEM. FTSGs, full-thickness
skin grafts; STSGs, split-thickness skin grafts.
C
Capillary density (cm/cm2)
400.00
350.00
300.00
250.00
200.00
150.00
fresh skin
100.00
3d old skin
50.00
7d old skin
0.00
0
Capillary diameter (um)
B
2
5
7
10
Days after Tx
25.00
Angiogenic wound bed reaction (bud
n/mm2)
A
25
fresh skin
3d old skin
20
7d old skin
15
10
5
0
0
3
5
7
10
Days after Tx
Fresh skin
3d old skin
20.00
7d old skin
15.00
10.00
5.00
0.00
0
3
5
7
Days after Tx
10
Figure 3. Human split-thickness skin grafts (0, 3, and 7 days of preservation) transplanted into an SCID mouse. Capillary
density (A) and capillary diameter (B) in wound bed. Angiogenic response of the host (C). Means ± SEM.
Copyright © American Burn Association. Unauthorized reproduction of this article is prohibited.
Journal of Burn Care & Research
Month/XXX 2013
6 Knapik et al
A
12
Total graft score
10
*
8
fresh skin
6
3d old skin
7d old skin
4
2
0
0
10
Days after Tx
B
Fresh
STSG
7d old
STSG
Transplantation
Figure 4. Histological evaluation of fresh and stored human STSGs 10 days after transplantation. A, Blinded pathological
skin quality evaluation. *P ≤.05 day 0 vs day 10. B, Representative hematoxylin and eosin sections of fresh and preserved
grafts before and 10 days after transplantation. STSGs, split-thickness skin grafts.
The quality of the graft, scored by a dermatopathologist in a blinded fashion 10 days after graft placement, showed a constant and nearly linear decrease
for all three storage periods starting from 10 points
at the time of grafting (completely viable skin, 100%)
and ending at 7.6 (24% decrease) for fresh skin, 6.4
(36% decrease) for 3-day preserved skin and 5.5
(45% decrease) for 7-day preserved skin, 10 days
after grafting. However, values were only significant
in the 7-day storage group.
Influence of Preservation on Skin Graft
Capillaries and Vascularization From the
Wound Bed (Mouse Vasculature)
The skin graft vasculature behaved in a similar manner in freshly transplanted and preserved grafts,
exhibiting a decrease in the number of present vessels of 67.4% in directly grafted skin and 61.9% in
7-day preserved skin at day 10 (Figure 5A and B).
The ingrowth of murine host capillaries was not
influenced by the storage time of the graft and was
slightly, but not significantly, higher in 7-day preserved skin samples when compared with freshly
grafted skin (Figure 5C). In both cases murine
capillaries were found in upper layers of the dermis in the graft (Figure 5D). Thus, replacement of
host capillaries occurred in a similar manner after
storage when compared with freshly transplanted
skin grafts.
Proliferation and Apoptosis Rates of
Preserved Grafts After Transplantation
Proliferation and apoptosis rates were determined
by immunohistochemistry in three separate layers:
muscle tissue of the wound bed, the lower dermis,
on the border of wound bed and graft, and finally in
the upper dermis, periphery of the graft. Our results
show that the proliferation rate measured on day 10
after graft placement remains constant in the periphery, middle part, and wound bed in all three tested
Copyright © American Burn Association. Unauthorized reproduction of this article is prohibited.
Journal of Burn Care & Research
Volume XXX, Number XXX
C
20.00
Fresh skin
7d old skin
*
18.00
16.00
*
14.00
12.00
10.00
8.00
6.00
4.00
2.00
0.00
0
Days after Tx
20.00
Number of mouse capillaries/0.5mm2
Number of human capillaries/0.5mm2
A
Knapik et al 7
18.00
Fresh skin
7d old skin
16.00
14.00
12.00
10.00
8.00
6.00
4.00
2.00
0.00
10
10 days after Tx
B
D
Epidermis
Human STSG
Fresh
STSG
Fresh
STSG
Host wound bed
7d old
STSG
7d old
STSG
Human STSG
Host wound bed
0
10
Figure 5. Evaluation of capillary density in STSGs 10 days after transplantation (A) human capillaries. B, Representative
slides of human CD31 graft vasculature of fresh and preserved human STSGs before and 10 days after grafting. C, Mouse
capillaries. D, Representative slides of mouse CD31 positive vasculature in fresh and preserved human STSGs 10 days after
graft placement. *P ≤ .05 day 0 vs day 10. STSGs, split-thickness skin grafts.
groups (Figure 6A). The preservation time of the
graft did not influence the proliferation rate. Furthermore, we observed differences in the proliferation rate between the three defined layers: the upper
dermis revealed ~×4 higher proliferation rates than
the middle dermal part of the graft or the wound
bed itself. The proliferation rate ranged between <1
and <8% depending on the layer, reaching the highest level in the upper part of the dermis. The rate of
apoptotic cells in general remained on a low level.
The results did not show significant differences
among the investigated skin layers.
Modern plastic and reconstructive surgery has to face
and integrate the ongoing developments in tissue
engineering in order to increase early wound coverage and reduce patient morbidity. This is particularly
of importance for the treatment of severely burned
patients, where often not enough autologous donor
skin is available for transplantation and rapid sufficient defect coverage is urgently needed to increase
patient survival.16 In this situation, often only skin
allografts or xenografts are available for temporary
B
12.00%
Upper dermis
Middle Dermis
Muscle
10.00%
8.00%
6.00%
4.00%
2.00%
0.00%
0
3
Skin storage time (days)
7
Apoptosis (%) 10days after grafting
Proliferation rate (%) 10 days after grafting
A
DISCUSSION
8.00%
Upper dermis
Middle Dermis
Muscle
6.00%
4.00%
2.00%
0.00%
0
3
Skin storage time (days)
7
Figure 6. Proliferation (A) and apoptosis rate (B) of cells in wound bed (muscle) and grafts (upper and lower dermis) 10
days after grafting.
Copyright © American Burn Association. Unauthorized reproduction of this article is prohibited.
8 Knapik et al
coverage followed by a long recovery period, making
use of the small amount of available skin by expansion and growth of keratinocyte sheets.17 Still, wound
coverage and eventually skin quality after these procedures are far from satisfactory, resulting in numerous reoperations.18 In recent times medical centers
also need to fulfill the requirements of “tissue directives” (2004/23/EC, 2006/17/EC, 2006/86/
EC), European regulations (2007/1394/EC), and
local/national legislation.19 Olender et al18 pointed
out that meeting the criteria set by law leads to
considerable financial expenses to adapt facilities
and train personnel. This in fact could become a
limiting factor for small hospitals. However, many
tissue banks have been involved in the procedure
and directive development process and are already
accordingly prepared. In addition, practitioners of
young disciplines like tissue engineering, allograft
revitalization, or stem cell–based therapies are fully
aware of the legal regulations for future product
requirements.20,21
Full-thickness skin substitutes were introduced
into the clinic with the tissue-engineering era.22–24
However, none of these dermo-epidermal substitutes were able to overcome the problem resulting
from the requirements of an FTSG to rapidly recruit
adequate oxygen and nutrient supply. This issue
often leads to total or partial tissue necrosis.25–27
In a previous experimental study we observed
that the integrity and composition of preserved
human skin grafts12 did not show deterioration after
a storage period of up to 7 days in moist gauze at
4 to 6°C. Cell survival was equal in fresh and preserved grafts, but the metabolic rate was decreased
by ~50% by day 3 after storage, indicating that cooling of the skin grafts reduced metabolism, leading to
lower energy demands and a decreased susceptibility to lack of oxygen. This state could be regarded
as a torpor-like state, as often seen in nature during
hibernation.14 Cell division and migration, as well
as protein translation and transcription, are arrested
during torpor.28,29
In light of these findings it was the goal of this
study to investigate the influence of human skin graft
storage up to 7 days at 4 to 6°C and the reaction of
the wound bed, in particular the angiogenic response
in vivo. In the first step we compared the revascularization process of human FTSGs with that of
human STSGs. Intravital microscopy results showed
no significant differences with respect to angiogenic
response and vascularity, even though there was a
trend toward an earlier peak in wound bed angiogenesis in STSGs. This is in line with the observation of
our previous studies investigating mouse FTSGs.15,30
Journal of Burn Care & Research
Month/XXX 2013
However, capillary widening of the wound bed was
much less pronounced after human skin graft application, when compared with the mouse model. This
could at least in part also be attributed to the different mouse species (SCID) studied. The fact that
there merely was a trend toward a different timing
in capillary widening between FTSGs and STSGs
may be accounted for by the small number of experiments. To our knowledge, no differences in the
wound bed reaction between STSGs and FTSGs
have been described before.
Preservation of human STSGs up to 7 days did
not influence the vascular response of the wound
bed in vivo. Typical angiogenic changes such as
capillary widening, increase of capillary density,
and capillary bud formation, occurred independent
of storage time in STSGs of all groups exhibiting
the same timing and reaching similar magnitudes.
From this it can be derived that the expression of
angiogenic factors during storage most likely is not
significantly increased indicating hypoxia-tolerance mechanisms within the skin grafts yet to be
defined. Hypoxia and associated factors like HIF1α are highly capable of inducing angiogenesis.31,32
In a previous study we were able to show, that the
hypoxic skin graft conveys a strong angiogenic
effect onto the wound bed by up-regulating HIF1α and subsequently VEGF peaking within the first
72 hours and leading to sprouting angiogenesis.30
Because the angiogenic response of the wound
was not different after skin graft storage it can be
deducted that most likely HIF-1α expression was
not significantly different, hinting at a similar level
of tissue hypoxia. However, further studies are
needed to investigate this hypothesis.
Next to this, the present study illustrates that
the preservation of skin grafts at 4 to 6°C for 7
days and subsequent transplantation into a warm
organism resulting in rewarming of the STSG lead
to significant changes in graft score and quality
after 7 days of storage at 4 to 6°C. Of interest,
histological graft score after storage of 7 days only,
that is, without transplantation into the skinfold
chamber, was not diminished.12 Thus, transplantation of the grafts into the warm mouse organism
revealed potential damaging factors in vivo, which
were not seen after storage and histological evaluation alone. This again points toward the hypoxiaprotective mechanisms with reduced metabolism
during storage.
Vascular ingrowth of wound bed vessels with subsequent partial replacement of existing graft vasculature has been shown to be a crucial step in skin
graft healing and survival.33,34 In this study STSG
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Journal of Burn Care & Research
Volume XXX, Number XXX
vasculature degeneration as well as vessel ingrowth
from the mouse wound bed was similar for freshly
harvested and 7-day stored skin grafts. This once
again suggests a similar hypoxia-level within the skin
graft before and after 7 days of storage. However,
autochthonous human graft vessel count within
the STSGs decreased significantly both for freshly
transplanted and 7-day stored skin grafts, confirming vessel degeneration and replacement with host
mouse vasculature over time. Mouse vessels were
detected in the upper dermis of the human skin
graft, depicting the complete vessel ingrowth from
the wound bed.
Last, immunohistological studies showed that
proliferation and apoptosis in STSGs did not reveal
significant differences after skin graft storage up to 7
days. Both rates were generally low and below 10%
in all study groups. The phenomenon that cooling slows down degenerative processes and oxygen
consumption is a well-known fact applied in clinical routine, for example, in heart surgery or neurosurgery.35,36 Our study and previous experiments
suggest, that human skin grafts stored under severe
conditions (4–6°C, up to 7 days without oxygen
and nutrient supply) are not damaged and most
likely equally hypoxic as fresh skin grafts. However,
skin graft quality is slightly decreased after 7 days of
storage and transplantation into a living organism.
This suggests that even when a graft microscopically
appears viable and without change in tissue composition after preservation, still underlying cell damage
may be present. The actual performance of the skin
graft in fact may only be revealed after transplantation into the living organism. We have shown in
a previous study that cell viability of the skin graft
dropped within the first 3 days of storage significantly to approximately 52%, reaching finally a value
of 44.1% in comparison with fresh skin until 14 days
of storage.12 This may in fact influence graft performance after transplantation and reveals an underlying tissue damage of the graft, if it is stored at 4 to
6°C in saline gauze. In accordance with solid organ
preservation special nutrient media or conditions
(eg, University of Wisconsin solution, histidine-tryptophan-ketoglutarate solution, Dulbecco’s Modified
Eagle Medium, cryopreservation, etc)9,37 should
be evaluated in this context to improve graft performance in vivo after storage. On the basis of this
evaluation studies of new storage media also should
be verified in an in vivo approach. Furthermore, the
findings of the current study could be translated and
used to design a new approach in skin tissue engineering by expression of hypoxia-tolerance–inducing
factors in full-thickness skin substitutes.
Knapik et al 9
ACKNOWLEDGMENTS
This study was financially supported by the Swiss National
Science Foundation (SNF-Grant 310030_127366). Drawings for Figure 1 were kindly provided by Stefan Schwyter,
Technical Drawer, Graphics Department, University Hospital Zürich, Zürich, Switzerland.
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