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Year : 2019  |  Volume : 27  |  Issue : 2  |  Page : 49-55

The effects of different amounts of thrombin application on fat graft viability in rats: An experimental study

1 Plastic Reconstructive and Aesthetic Surgery Clinic, Dr. Lütfi Kirdar Kartal Training and Research Hospital, Istanbul, Turkey
2 Plastic Reconstructive and Aesthetic Surgery Clinic, Bagcilar Training and Research Hospital, Istanbul, Turkey
3 Department of Pathology, Marmara University Faculty of Medicine, Istanbul, Turkey

Date of Web Publication27-Mar-2019

Correspondence Address:
Dr. Emre Guvercin
Plastic Reconstructive and Aesthetic Surgery Clinic, Dr. Lütfi Kirdar Kartal Training and Research Hospital, Istanbul
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Source of Support: None, Conflict of Interest: None

DOI: 10.4103/tjps.tjps_53_18

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Aims: The most important disadvantage of fat graft, which is also a late-term complication, is graft resorption. The aim of this study is to evaluate the effect of thrombin, which is reported to increase the tissue regeneration and angiogenesis in many areas, to viability of fat graft. Settings and Design: Twenty Wistar-Albino type adult male rats were used in the study. They were divided into four groups as one control group and three experimental group. Subjects and Methods: Inguinal fat pads were excised and reduced to 500 mg (±1 mg) in all animals. To obtain thrombin, 3 animals were sacrificed. One percent amount of 0.25 mg lidocaine hydrochloride was injected to the scapular regions of the animals. Afterward, subcutaneous cavities were formed there to place the fat tissue inside. After placing the graft, varying proportions of thrombin were injected to the animals in the experimental groups. No thrombin was not injected to the control group. After 90 days, the experimental animals were sacrificed, and the fat grafts were removed. Statistical Analysis Used: The data obtained from control and experimental groups were analyzed using SPSS software (version 20.0; SPSS Inc., Chicago, IL, USA). Normal fat ratio, cyst or vacuole development, inflammation, fibrosis, microvascular density, apoptosis, and weight score differences between groups were compared using Kruskal–Wallis test. To compare two groups with different scores, Mann–Whitney U test was used. The statistical significance level was accepted as 0.05 (P = 0.05). Results: Macroscopic, histological, and statistical evaluations showed that thrombin has reduced the weight and volume loss on fat graft, increased viable fat cell amount and reduced inflammation on receptive area. Conclusion: The positive effects of thrombin on the viability of fat graft have given us courage to use it in further studies. Longer follow-ups are necessary and more studies are required to use it in clinical practice in combination with fat grafts.

Keywords: Autologous tissue transfer, fat grafts, graft viability, thrombin

How to cite this article:
Kacmaz C, Gideroglu K, Guvercin E, Filinte GT, Bozkurt M, Filinte D. The effects of different amounts of thrombin application on fat graft viability in rats: An experimental study. Turk J Plast Surg 2019;27:49-55

How to cite this URL:
Kacmaz C, Gideroglu K, Guvercin E, Filinte GT, Bozkurt M, Filinte D. The effects of different amounts of thrombin application on fat graft viability in rats: An experimental study. Turk J Plast Surg [serial online] 2019 [cited 2022 Oct 5];27:49-55. Available from: http://www.turkjplastsurg.org/text.asp?2019/27/2/49/255007

  Introduction Top

Autologous fat grafting is a popular procedure that has been performed at an increasing rate in recent years for both aesthetic and reconstructive purposes.[1] There are advantages such as the trustee of autologous fat tissue, a wide donor area, and practical application; the most important disadvantage and a late complication is graft resorption.[2],[3]

Many studies have been done regarding the reduction of fat graft loss; however, the clinical implications of these studies are not at the expected level.[4] Many experimental studies have also been performed exploring means of reducing fat graft absorption.[5],[6],[7] Lipoaspiration with low negative pressure vacuums to reduce mechanical trauma on fat graft, adding vitamin E and steroids to support microenvironment and removing inflammatory mediators with irrigation are some examples of techniques on these studies.[6],[7]

According to some studies, fat grafting to well-vascularized areas improves fat graft uptake, and it is thought that this increase is due to high vascular density.[8] Various methods have been tested to increase vascular density on the recipient area, and some have been shown to be experimentally effective.[7],[9] Fat graft loss in early stages is linked to necrosis due to acute ischemia and on to apoptosis in later stages.[10] This explains graft loss after neovascularization of graft tissue.[3]

Many growth factors in α-granules of platelets are known to stimulate cell proliferation and regeneration.[11] To take advantage of this effect of platelets, the use of platelet-rich plasma (PRP) with fat grafts has become very popular in recent years; as platelet count may be up to 1 million/mm 3 in PRP.[12]

Many substances secreted from platelets are angiogenic and stimulate tissue regeneration. The aim of this study was to evaluate the effect of thrombin, known to be such a stimulant for platelets on viability of transfer when applied to recipient site of graft.

  Subjects and Methods Top

The study was performed in a university hospital, center for animal research; and histopathological studies were performed in the pathology department. The University Ethical Committee for animal experiments and research certified the study. Twenty Wistar adult male albino rats weighing 250–300 g were housed in a room at temperature of 20–24°C with, 40%–60% relative humidity and 12-h light cycle before the study. Each subject was kept in individual metal cages after the operation. Subjects were separated into four groups: three experimental and one control group. Three animals were euthanized to obtain thrombin from whole body blood.

The groups were defined as follows:

  1. Experimental group: 500 mg fat tissue; 0.60 cc thrombin combined Group (1\2)
  2. Experimental group: 500 mg fat tissue; 0.30 cc thrombin combined Group (1\1)
  3. Experimental group: 500 mg fat tissue; 0.15 cc thrombin combined Group (1\0.5)
  4. Control group: Only 500 mg fat tissue; without thrombin combined Group (1\0).

All animals received 100 mg/kg ketamine and 10 mg/kg xylazine intraperitoneally as anesthesia and 10 mg/kg cefazolin was administered intramuscularly to prevent infection.

Obtaining the fat grafts

The inguinal region of the subjects was shaved, and a line from penis to hip joint was marked for skin incision. In order to achieve additional local anesthesia, 0.25 cc of lidocaine hydrochloride (1% concentration) was injected at incision line. Surgical field was cleaned with chlorhexidine solution. Inguinal fat pads were excised through the incision and subsequently reduced to pieces weighing 500 mg (±1 mg). Fat tissues were washed with 0.9% saline solution. Inguinal incisions were sutured with 4–0 silk [Figure 1].
Figure 1: Inguinal fat pads were excised and reduced to 500 mg (±1 mg)

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Obtaining the thrombin

GLO® PRP (GloFinn Oy, Salo, Finland) kit was used to obtain thrombin from venous blood. Three rats were euthanized to obtain whole body blood, and thoracotomy was performed to obtain 19.5 cc of blood directly from the right ventricle. Then, 1 cc anticoagulant citrate dextrose-A and 9 cc blood were drawn into a syringe to create 10 cc mixture; 8 cc of this mixture was transferred from syringe to empty test tube. To activate the 8 cc mixture, 1.7 cc of calcium gluconate solution was added. The same process was repeated to obtain thrombin from the remaining blood. Tubes were kept upright at room temperature of 20–24°C for 45 min before centrifuged at 3000 rpm for 8 min. Total of approximately 6.5 cc thrombin was obtained [Figure 2].
Figure 2: Thrombin (below) obtained from whole body blood (above) of 3 rats

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Adapting the fat grafts and thrombin to recipient area

To achieve local anesthesia, 0.25 cc of lidocaine hydrochloride 1% concentration was injected to scapular region of the animals. Subcutaneous pockets were created in the area for fat graft insertion. Scapular region was chosen because there is almost no fat tissue. After fat graft insertion to recipient site, thrombin was injected around the fat grafts in different quantities for each experimental group; no injection was performed on control group. Incisions were closed with 4–0 silk sutures [Figure 3].
Figure 3: Placement of fat grafts in subcutaneous scapular cavity (left) and thrombin injection to fat graft in experiment groups (right)

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Betadine solution was used for wound care. At 90th day, animals were euthanized, fat grafts were taken from recipient site, washed with 0.9% saline solution, weighed with precision weighing instruments, and results were recorded for each subject. All samples were fixed with 10% formalin solution for histopathological study [Figure 4].
Figure 4: Dissection and removal of fat grafts after 90 days

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Histological assessment

Fat grafts obtained from the 20 subjects of control and experimental groups were fixed with 10% formalin solution. After routine tissue follow-up, samples were embedded in paraffin. Sections 4 μm in size were taken from the paraffin blocks and stained with Masson's trichrome and h and e stains. Samples were evaluated under light microscope. Normal fat ratio, cyste or vacuole development, inflammation, fibrosis, microvascular density, and apoptosis were studied. Normal fat ratio, cyst or vacuole development, inflammation, and fibrosis were scored on scale of 0–3 (0: None, 1: Mild, 2: Moderate, 3: High). For microvascular density and apoptosis, vascular structures, and apoptotic cells were counted in 10 high power fields, ×400).

Tissue apoptosis was demonstrated using terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling method and ApopTag® Plus Peroxidase in situ Apoptosis Detection Kit (S7101,; Millipore Corp., Bedford, MA, USA). Sections of 4 mm thickness were taken from fat grafts with microtome, deparaffinized for 1 night at 37°C and hydrated using xylene and alcohol series. Sections were then incubated with Proteinase K (20 μg/cc) for 15 min at about room temperature. Endogenous peroxidase activity in tissues was suppressed with 3% hydrogen peroxide solution (pH 7.4 in phosphate buffered saline [PBS]) for 5 min period. The sections were washed with PBS, then incubated in equilibration buffer for 30 min at room temperature, and for 60 min at 37°C with working strength terminal deoxynucleotidyl transferase enzyme solution. Slides were washed with working strength stop/wash buffer for 10 min. Anti-digoxigenin-conjugate was applied to sections at room temperature, and sections were incubated again for 30 min. 3,3'Diaminobenzidine (Peroxidase Substrate) was applied to the sections. They were washed with PBS and incubated for 5 min. After washing with distilled water, they were stained with Mayer's hematoxylin. The sections were dehydrated with alcohol and then evaluated.

Statistical analysis

The data obtained from control and experimental groups were analyzed using SPSS software (version 20.0; SPSS Inc., Chicago, IL, USA). Normal fat ratio, cyst or vacuole development, inflammation, fibrosis, microvascular density, apoptosis, and weight score differences between groups were compared using Kruskal–Wallis test. To compare two groups with different scores, Mann–Whitney U test was used. The statistical significance level was accepted as 0.05 (P = 0.05).

  Results Top

Macroscopic findings

After all subjects were euthanized, incisions were made at recipient site, dissection was performed, and fat grafts were accessed through panniculus tissue. Tissue surrounding fat grafts in control group was found to be fibrotic; grafts were attached to the surrounding tissue and were quite hard. In experimental groups, fat grafts had only loose connective tissue attaching them to surrounding tissue, were dissected easily, and were quite soft. In addition, the volume of the fat grafts in experimental groups was greater than in the control group.

Weight measurement

The average weight of the fat graft in groups injected with 0.15 cc and 0.60 cc of thrombin was significantly higher than that of control group (P = 0.040 and 0.048, respectively).

There was no difference in average weight of graft between control group and 0.30 cc thrombin group or; between thrombin groups [Table 1] and [Graphic 1].
Table 1: Comparison of All Groups with Kruskal-Wallis Test

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Histological findings

Normal fat ratio was significantly higher in 0.30 cc thrombin group than control group (P = 0.032). There was no significant difference between 0.15 cc and 0.60 cc thrombin groups and control group. There was no significant difference between 0.15 cc, 0.60cc, and 0.30 cc thrombin groups [Table 1], [Graphic 2] and [Figure 5].

Figure 5: Normal fat cells from 0.60 cc thrombin group, shown with H and E staining

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Inflammation was significantly lower in 0.30cc and 0.60 cc thrombin groups compared to control group (P = 0.032, P = 0.024, respectively) and in 0.15 cc thrombin group, inflammation was lower than that found in to 0.60 cc thrombin group (P = 0.048). There were no other significant differences between other matches [Table 1], [Graphic 3] and [Figure 6].

Figure 6: (a and b) Inflammation and cystic degeneration in control group with H and E staining. (c) Severe fibrosis, inflammation, and cystic degeneration with masson trichrome stain in control group

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Microvascular density was higher in thrombin groups compared to control group, but not statistically significant (P > 0.05). There was no significant difference in fibrosis or cyst formation between groups (P > 0.05). Apoptosis occurred less in thrombin groups compared to control group, but the difference was not statistically significant (P > 0.05) [Table 2] and [Figure 7].
Table 2: Apoptosis Distribution According to Groups

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Figure 7: (a) Inflammation and apoptotic cells in a fibrotic ground, in control group stained with terminal deoxynucleotidyl transferase dUTP nick end labeling method. (b) Fewer apoptotic fat cells seen in 30 cc thrombin group

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  Discussion Top

Soft-tissue defects can occur many reasons, including burns, tumor surgery, trauma or congenital anomalies, and they play a large role in plastic surgery. Alternative methods such as local and free flaps, dermal fat grafts, synthetic materials, and autologous fat grafts have been developed to reconstruct soft-tissue defects.[13] The most common cause of dissatisfaction after reconstructive surgery is lack of sufficient subcutaneous fat tissue.[14] Recently, as the use of minimally invasive methods grew, injectable materials have been used for soft-tissue defect repair.[1] Although there is a wide variety of synthetic fillers, there is no ideal material that meets all expectations.[15] Unnatural result, foreign body reaction, and lack of permanent effect are some examples of disadvantages of these materials. In addition, these products are very expensive.[16]

One of the basic principles of plastic surgery is “to replace like with like.”[17],[18] Autologous tissue has still been accepted as the most ideal reconstruction materials.[15] The most important advantages of fat grafting are being able to fill large soft-tissue defects, no risk of allergic reaction, wide donor site, good tissue conformity, and procedure is simple to perform with low morbidity.[19],[20] Autologous fat grafting also has disadvantages. Necrosis, infection, prolonged edema, cyst formation, atrophy, irregularity, and impermanence are some examples of possible negative results.[2],[3],[21] Outcomes of autologous fat grafting are unpredictable. Volume loss of 20%–50% may occur in long-term.[2],[3],[4],[22] Consensus has not yet been achieved on optimal technique, persistence of grafts, or other related issues.[2] The common idea is there has always been graft loss but in varying amounts.[1] Factors being examined that have an impact on fat grafting success include:[23],[24]

  • Effect of anesthesia
  • Method for taking fat grafts with vacuum or excision
  • Effect of cannula when taking graft and injecting fat
  • Removal of blood from grafts-purification
  • Traumatization of fat grafts during purification
  • Contact with air
  • Cryopreservation of fat cells
  • Contamination
  • Vascularization of recipient site.

Many studies have been performed to reduce fat graft loss and to improve fat graft viability.[25],[26],[27] Parallel to these studies, it has also been was shown that fat cells have low tolerance to ischemia.[28],[29] Placing fat grafts into a well-vascularized area has been shown to improve graft viability.[28],[30] Heimburg demonstrated that expansion of fat graft increases vascularization and resistance to ischemia.[29] In addition to injection of fat grafts to well-vascularized fields such as muscle tissue to improve viability, the consequences of increasing the vascularization of recipient site have also been investigated. Baran et al. showed that the viability of fat cells improved when they were injected into well-vascularized capsule tissue that occurred with a silicone block inserted to rats.[7]

During fat grafting, circulation of fat cells worsens, and they face partial trauma. Macrophages, histiocytes, and multinucleated giant cells migrate to recipient site, neutrophil infiltration occurs, and cellular debris is cleared.[31] Neovascularization starts on fat grafts 2–4 days after the transfer.[3],[32] This neovascularization is limited to peripheral cells. Fat cells continue to die until new vascular network reaches a sufficient level. Fibrous tissue increases in later stages but is insufficient to complete volume lost.[31] To ensure proper tissue perfusion of fat cells, oxygen, and other needs of these cells must be met.[32],[33] The viability of adipocytes in autologous fat grafts depends on early neovascularization.[33]

In recent studies, it has been reported that functional vessel density reaches to measurable level on the 15th day and continues to increase until 60th day; however, it then decreases by 50% in 90–120 days.[32] Nishimura et al. reported that the reduction of fat cell count continues after neovascularization occurs. Loss of fat cells in early period of transfer is due to acute necrosis; in later period, it is the result of apoptosis. This is explanation for ongoing fat graft loss after neovascularization.[3]

Thrombin (MW 33,700) is an unstable, protein-structured enzyme, formed by cleavage of plasma protein prothrombin. It has 2 independent receptors, 1 proteolytic (PAR1-PAR4) and 1 nonproteolytic receptor.[34] Many events induced by enzymatic reactions such as coagulation occurs through proteolytic receptors of thrombin.[35] Nonproteolytic thrombin receptors exhibit on many cells's surface and have a high affinity for thrombin; these effects are dose-dependent because of the receptor saturation.[36] Thrombin has been reported to affect inflammatory cells and stimulate immune cells by regulating the secretion of inflammatory cytokines.[37] It also stimulates monocytes, macrophage chemotaxis, fibroblastic cells, and endothelial cells, and plays an effective role in cell regeneration and wound healing.[38]

Although many studies have been performed to increase the fat graft viability, there has been none thus far that explored the effect of thrombin. Autologous thrombin was selected for use in the present study due to low cost, ease of application, and use of subject own blood. Thrombin stimulates secretion of many growth factors by binding its receptors to platelets.[12] We believe that the stimulatory effect of thrombin on cells occurs as a result of this mechanism.

In the study, based on macroscopic, histological, and statistical evaluation of parameters we examined, we found that weight loss, and thus loss of fat graft, decreased in thrombin-treated groups. Histological examination showed increase in viable fat cells in thrombin-treated groups, and this supports our hypothesis that thrombin reduces graft loss. Although microvascular density increased in thrombin groups when evaluated histologically, there was no statistically significant difference between control and experiment groups; we think that statistically significant results may be obtained by increasing the number of subjects in the study.

Significant decrease of inflammation is 1 of the parameters indicating the increasing effect of thrombin on graft viability because biological mediators secreted to the areas where inflammatory cells are concentrated stimulate migration of inflammatory cells to these areas. Inflammatory cells give harm to healthy cells by secreting various cytokines and increasing lysosomal enzymes and free radicals in the field, inhibiting the normal cycle of cells.[39] There was less adherence of fat cells to surrounding tissues seen on excision of fat grafts in thrombin groups, which also proves less inflammation occurred in those groups compared to control group. In the present study, we observed that different quantity of thrombin had different effects on fat graft weight, normal fat ratio, and inflammation.

Accordingly, we think that further study should be conducted with a better dose titration design to find optimum dose. Fat cell apoptosis was lower in thrombin-treated group than control group, but it was not significant ([P = 0.317 for 0.15 cc thrombin, P = 0.087 for 0.30 cc thrombin, and P = 0.111 for 0.60 cc thrombin] [Table 2]). The dose of thrombin between 0.15 and 0.30 cc may decrease apoptosis in fat grafts because of aforementioned dose-dependent properties of thrombin.

Scar formation and adhesions following skin grafting on mobile structures such as tendons and joints when treating trauma and burn injuries is a common problem.[40] Fat grafting is among treatment choices in these instances. The effect of thrombin found in the present study suggests that it could be effective at improving functional loss of mobile tissue under skin-grafted areas during contour restoration with fat grafting.

  Conclusion Top

We found that thrombin reduces fat graft weight loss by decreasing the inflammation level. According to the literature, so far it has not been used to improve fat graft uptake and reduce fat graft loss. We believe better dose-titrated studies should be done to find appropriate dose experimentally and that further clinical studies should be done to determine effects on human fat grafting procedures.

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Conflicts of interest

There are no conflicts of interest.

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  [Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5], [Figure 6], [Figure 7]

  [Table 1], [Table 2]


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