Plastic and Reconstructive Surgery • February 2015 Reply: Studies in Fat Grafting: Part I. Effects of Injection Technique on In Vitro Fat Viability and In Vivo Volume Retention; and Studies in Fat Grafting: Part II. Effects of Injection Mechanics on Material Properties of Fat Sir:
W
e would like to thank Dr. Jia et al. for their comments concerning our two reports, “Studies in Fat Grafting: Part I. Effects of Injection Technique on In Vitro Fat Viability and In Vivo Volume Retention” and “Studies in Fat Grafting: Part II. Effects of Injection Mechanics on Material Properties of Fat.” They first raise concerns regarding our use of the phrase “modified Coleman technique” and methods that do not strictly adhere to those described by Sydney Coleman encompassing gentle manual harvest, centrifugation refinement, and slow low-volume retrograde placement of fat. We agree that our methods do not directly mirror those of Dr. Coleman, and thus consciously referred to our techniques for fat harvest and injection as “modified.” Furthermore, the field of fat grafting is extremely heterogeneous, and although some studies have used centrifugation for processing, many others have relied on gravity separation or filtration to isolate fat for injection.1–4 Even when “Coleman” techniques are described in the literature, there can be variability in the centrifugation speed/force and time reported.5 Dr. Jia and colleagues also note that transfers between our “modified Coleman” control group and the Adipose Tissue Injector (TauTona Group, Menlo Park, Calif.) group were not identical, with the control group fat passing through a 60-cc syringe, to a 10-cc syringe, and then to 1-cc syringe for injection. In contrast, the Adipose Tissue Injector group obviates much of these transfers, as the 60-cc syringe was loaded directly onto the Adipose Tissue Injector device. This difference is, in fact, one of the key advantages to using the Adipose Tissue Injector device over a modified Coleman approach, as the Adipose Tissue Injector device minimizes the preinjection handling. Our two reports, “Studies in Fat Grafting: Part I” and “Studies in Fat Grafting: Part II,” were designed to prove this very point, that additional preinjection steps commonly used in various “Coleman” protocols are detrimental to fat. By avoiding multiple fat transfers, the Adipose Tissue Injector device preserves viability and avoids a “shear storm” fat would experience using the Coleman technique. An issue over the rate of injection in our studies is also brought forth, one which is confusing to us. As clearly stated in our second report, the Adipose Tissue Injector device delivers fat at a controlled flow rate of 0.5 cc/second. This flow rate can be set irrespective of actual volume delivered. As Dr. Jia et al. state, Coleman emphasizes retrograde delivery with small volumes (1/10 to 1/30 ml) per withdrawal. How this actually translates into flow rate is entirely dependent on time spent during withdrawal. Furthermore, in actual practice, although withdrawal of the syringe may realistically
448e
occur over 0.5 to 2 seconds, actual duration of injection may be far shorter, depending on force applied to the plunger and whether the cannula is “plugged.” It is for these reasons that the Adipose Tissue Injector device is, again, more “fat friendly.” Concerns are also raised over statistical analysis. We agree with Dr. Jia et al. that repeated measures analysis of variance should have been used to evaluate the histology scores recorded for each independent investigator; however, rerunning our statistics with this approach did not change our overall findings, with healthy (p = 0.045) and injury (p = 0.013) scores still remaining below the p < 0.05 threshold as detailed in our article. In addition, differences in volume retention between our studies and a report coauthored by Sydney Coleman is brought up.6 Interestingly, the same disparities Dr. Jia and colleagues mention with that previous study, including donor site, techniques, and recipient site, were already discussed in one of our earlier reports documenting the advantages of our model; namely, use of the scalp recipient site and micro–computed tomographic imaging allows for measurement of in vivo fat graft retention without the need for animal sacrifice and explantation of fat.6,7 Finally, Dr. Jia et al. note that the test conditions reported in Part II of our studies were far different from what the graft experiences in a “grafting session” and that questions arise over clinical translation. Frankly, we find this concern surprising considering the context of our two reports presented in back-to-back fashion in the same issue. Clearly, Part II is a material science article evaluating properties of fat. These findings go hand-inhand with fat retention findings in Part I showing superiority using the Adipose Tissue Injector device. Again, we would like to thank Dr. Jia and colleagues for their attention to our two studies in fat grafting. We agree that with the Adipose Tissue Injector device, fat experiences a lower shear environment, and along with savings in time and labor, more precision and reproducibility can be achieved to enhance fat graft outcomes. DOI: 10.1097/PRS.0000000000000978
Derrick C. Wan, M.D. Geoffrey C. Gurtner, M.D. Hagey Laboratory for Pediatric Regenerative Medicine
Michael T. Longaker, M.D., M.B.A. Hagey Laboratory for Pediatric Regenerative Medicine and Institute for Stem Cell Biology and Regenerative Medicine Stanford University Stanford, Calif. Correspondence to Dr. Wan Department of Surgery Child Health Research Institute Stanford University Medical Center 257 Campus Drive Stanford, Calif. 94305 [email protected]
Volume 135 Number 2 • Letters Correspondence to Dr. Longaker Institute for Stem Cell Biology and Regenerative Medicine Stanford University Medical Center 257 Campus Drive Stanford, Calif. 94305 [email protected]
DISCLOSURE Dr. Gurtner and Dr. Longaker have equity in the TauTona Group. ACKNOWLEDGMENTS This work was supported by National Institutes of Health grants R01 AG-25016, R01 DK-074095, and 1R01 HL104236-01 (to G.C.G.); the American College of Surgeons Franklin H. Martin Faculty Research Fellowship, the Hagey Laboratory for Pediatric Regenerative Medicine, and the Stanford University Child Health Research Institute Faculty Scholar Award (to D.C.W.); and National Institutes of Health grants R01 DE021683-01 and RC2 DE020771, the Oak Foundation, and the Hagey Laboratory for Pediatric Regenerative Medicine (to M.T.L.). REFERENCES 1. Jianhui Z, Chenggang Y, Binglun L, et al. Autologous fat graft and bone marrow-derived mesenchymal stem cells assisted fat graft for treatment of Parry-Romberg syndrome. Ann Plast Surg. 2014;73(Suppl 1):S99–S103. 2. Nguyen PS, Desouches C, Gay AM, Hautier A, Magalon G. Development of micro-injection as an innovative autologous fat graft technique: The use of adipose tissue as dermal filler. J Plast Reconstr Aesthet Surg. 2012;65:1692–1699. 3. Zhu M, Cohen SR, Hicok KC, et al. Comparison of three different fat graft preparation methods: Gravity separation, centrifugation, and simultaneous washing with filtration in a closed system. Plast Reconstr Surg. 2013;131:873–880. 4. Glashofer M, Lawrence N. Fat transplantation for treatment of the senescent face. Dermatol Ther. 2006;19:169–176. 5. Rohrich RJ, Sorokin ES, Brown SA. In search of improved fat transfer viability: A quantitative analysis of the role of centrifugation and harvest site. Plast Reconstr Surg. 2004;113:391–395; discussion 396–397. 6. Thanik VD, Chang CC, Lerman OZ, et al. A murine model for studying diffusely injected human fat. Plast Reconstr Surg. 2009;124:74–81. 7. Chung MT, Hyun JS, Lo DD, et al. Micro-computed tomography evaluation of human fat grafts in nude mice. Tissue Eng Part C Methods 2013;19:227–232.
The Vasculosome Theory Sir: read with great interest the article entitled “True and ‘Choke’ Anastomoses between Perforator Angiosomes: Part II. Dynamic Thermographic Identification,” by Chubb et al. Interestingly, the findings of this study could better be explained by the “perforasome theory” rather than the “angiosome theory.”1 Both theories provide a guide to identify tissue units that can be reliably transferred in a single stage, based on a single
I
source of blood supply. However, both theories cannot reliably predict the exact quantity of tissue that can be supported by a single feeding vessel.2 Taylor and Palmer defined the angiosome as a vascular territory, which is marked by the presence of reduced caliber vessels in its periphery.3 The caliber by definition is narrow enough not to allow independent perfusion of the adjacent angiosome. Eventually, Taylor et al. also defined a perforasome as an inherent part of the angiosome.4 In the current article, Chubb et al. investigate reduced caliber vessels connecting perforasomes.5 It seems to contradict their angiosome theory, because by definition there cannot be narrow caliber vessels within an angiosome. The only reasonable explanation to sustain the description by Taylor et al. of a perforasome being a part of the angiosome, is to concede that angiosomes are not defined territories by themselves but are territories defined by several perforasomes (i.e., ∑perforasomes = an angiosome). Saint-Cyr et al. state, “Each perforator has its own vascular territory, called a perforasome, which carries a multidirectional flow pattern that is highly variable and complex.”1 Each perforasome is connected to its adjacent perforasomes by means of linking vessels, which could be “large-caliber” or “narrow-caliber” channels. However, the direction in which a larger tissue territory could be harvested based on a single perforator cannot be predicted based on the perforasome theory. McGregor and Morgan defined an axial flap as “a single pedicled flap which has an anatomically recognized arteriovenous system running along its long axis.”6 Branches of adjacent perforators, connected together by large-caliber channels, can form such a vascular axis. A unit of tissue incorporating this vascular axis and nourished by a single “feeder vessel” or perforator could be considered to be a “vasculosome.” In other words, any feeder vessel arising from a source vessel and contributing to the “vascular axis” should be sufficient to support the entire vasculosome independently. In fact, the objective of “flap delay” is to induce the formation of a neovasculosome with a centrally placed “vascular axis.”7 For a cutaneous vasculosome, suprafascial directionality of a perforator points toward the large-caliber channels with which it forms a vascular axis in the suprafascial plane.8 A single perforator supplying this vascular axis should be adequate to support a flap including the two (or more) adjacent “skin-perforasomes,”9 which will constitute a vasculosome (Fig. 1). Chubb et al. have suggested the use of thermography for detecting this vascular axis to aid flap elevation.5 Indeed, accurate mapping of a particular cutaneous vascular axis will result in better delineation of the cutaneous vasculosome and thus elevation of a more reliable flap. The muscle vasculosome has a vascular axis in the muscle. A single feeding vessel supplying any part of the muscle can nourish the entire muscle, provided that a vascular axis exists within the muscle (Fig. 2). If there are “narrow-caliber” channels linking adjacent muscle units, reliable harvest of both territories based on a single feeder is unlikely.10 Even so-called traditional minor pedicles can
449e
W
e would like to thank Dr. Jia et al. for their comments concerning our two reports, “Studies in Fat Grafting: Part I. Effects of Injection Technique on In Vitro Fat Viability and In Vivo Volume Retention” and “Studies in Fat Grafting: Part II. Effects of Injection Mechanics on Material Properties of Fat.” They first raise concerns regarding our use of the phrase “modified Coleman technique” and methods that do not strictly adhere to those described by Sydney Coleman encompassing gentle manual harvest, centrifugation refinement, and slow low-volume retrograde placement of fat. We agree that our methods do not directly mirror those of Dr. Coleman, and thus consciously referred to our techniques for fat harvest and injection as “modified.” Furthermore, the field of fat grafting is extremely heterogeneous, and although some studies have used centrifugation for processing, many others have relied on gravity separation or filtration to isolate fat for injection.1–4 Even when “Coleman” techniques are described in the literature, there can be variability in the centrifugation speed/force and time reported.5 Dr. Jia and colleagues also note that transfers between our “modified Coleman” control group and the Adipose Tissue Injector (TauTona Group, Menlo Park, Calif.) group were not identical, with the control group fat passing through a 60-cc syringe, to a 10-cc syringe, and then to 1-cc syringe for injection. In contrast, the Adipose Tissue Injector group obviates much of these transfers, as the 60-cc syringe was loaded directly onto the Adipose Tissue Injector device. This difference is, in fact, one of the key advantages to using the Adipose Tissue Injector device over a modified Coleman approach, as the Adipose Tissue Injector device minimizes the preinjection handling. Our two reports, “Studies in Fat Grafting: Part I” and “Studies in Fat Grafting: Part II,” were designed to prove this very point, that additional preinjection steps commonly used in various “Coleman” protocols are detrimental to fat. By avoiding multiple fat transfers, the Adipose Tissue Injector device preserves viability and avoids a “shear storm” fat would experience using the Coleman technique. An issue over the rate of injection in our studies is also brought forth, one which is confusing to us. As clearly stated in our second report, the Adipose Tissue Injector device delivers fat at a controlled flow rate of 0.5 cc/second. This flow rate can be set irrespective of actual volume delivered. As Dr. Jia et al. state, Coleman emphasizes retrograde delivery with small volumes (1/10 to 1/30 ml) per withdrawal. How this actually translates into flow rate is entirely dependent on time spent during withdrawal. Furthermore, in actual practice, although withdrawal of the syringe may realistically
448e
occur over 0.5 to 2 seconds, actual duration of injection may be far shorter, depending on force applied to the plunger and whether the cannula is “plugged.” It is for these reasons that the Adipose Tissue Injector device is, again, more “fat friendly.” Concerns are also raised over statistical analysis. We agree with Dr. Jia et al. that repeated measures analysis of variance should have been used to evaluate the histology scores recorded for each independent investigator; however, rerunning our statistics with this approach did not change our overall findings, with healthy (p = 0.045) and injury (p = 0.013) scores still remaining below the p < 0.05 threshold as detailed in our article. In addition, differences in volume retention between our studies and a report coauthored by Sydney Coleman is brought up.6 Interestingly, the same disparities Dr. Jia and colleagues mention with that previous study, including donor site, techniques, and recipient site, were already discussed in one of our earlier reports documenting the advantages of our model; namely, use of the scalp recipient site and micro–computed tomographic imaging allows for measurement of in vivo fat graft retention without the need for animal sacrifice and explantation of fat.6,7 Finally, Dr. Jia et al. note that the test conditions reported in Part II of our studies were far different from what the graft experiences in a “grafting session” and that questions arise over clinical translation. Frankly, we find this concern surprising considering the context of our two reports presented in back-to-back fashion in the same issue. Clearly, Part II is a material science article evaluating properties of fat. These findings go hand-inhand with fat retention findings in Part I showing superiority using the Adipose Tissue Injector device. Again, we would like to thank Dr. Jia and colleagues for their attention to our two studies in fat grafting. We agree that with the Adipose Tissue Injector device, fat experiences a lower shear environment, and along with savings in time and labor, more precision and reproducibility can be achieved to enhance fat graft outcomes. DOI: 10.1097/PRS.0000000000000978
Derrick C. Wan, M.D. Geoffrey C. Gurtner, M.D. Hagey Laboratory for Pediatric Regenerative Medicine
Michael T. Longaker, M.D., M.B.A. Hagey Laboratory for Pediatric Regenerative Medicine and Institute for Stem Cell Biology and Regenerative Medicine Stanford University Stanford, Calif. Correspondence to Dr. Wan Department of Surgery Child Health Research Institute Stanford University Medical Center 257 Campus Drive Stanford, Calif. 94305 [email protected]
Volume 135 Number 2 • Letters Correspondence to Dr. Longaker Institute for Stem Cell Biology and Regenerative Medicine Stanford University Medical Center 257 Campus Drive Stanford, Calif. 94305 [email protected]
DISCLOSURE Dr. Gurtner and Dr. Longaker have equity in the TauTona Group. ACKNOWLEDGMENTS This work was supported by National Institutes of Health grants R01 AG-25016, R01 DK-074095, and 1R01 HL104236-01 (to G.C.G.); the American College of Surgeons Franklin H. Martin Faculty Research Fellowship, the Hagey Laboratory for Pediatric Regenerative Medicine, and the Stanford University Child Health Research Institute Faculty Scholar Award (to D.C.W.); and National Institutes of Health grants R01 DE021683-01 and RC2 DE020771, the Oak Foundation, and the Hagey Laboratory for Pediatric Regenerative Medicine (to M.T.L.). REFERENCES 1. Jianhui Z, Chenggang Y, Binglun L, et al. Autologous fat graft and bone marrow-derived mesenchymal stem cells assisted fat graft for treatment of Parry-Romberg syndrome. Ann Plast Surg. 2014;73(Suppl 1):S99–S103. 2. Nguyen PS, Desouches C, Gay AM, Hautier A, Magalon G. Development of micro-injection as an innovative autologous fat graft technique: The use of adipose tissue as dermal filler. J Plast Reconstr Aesthet Surg. 2012;65:1692–1699. 3. Zhu M, Cohen SR, Hicok KC, et al. Comparison of three different fat graft preparation methods: Gravity separation, centrifugation, and simultaneous washing with filtration in a closed system. Plast Reconstr Surg. 2013;131:873–880. 4. Glashofer M, Lawrence N. Fat transplantation for treatment of the senescent face. Dermatol Ther. 2006;19:169–176. 5. Rohrich RJ, Sorokin ES, Brown SA. In search of improved fat transfer viability: A quantitative analysis of the role of centrifugation and harvest site. Plast Reconstr Surg. 2004;113:391–395; discussion 396–397. 6. Thanik VD, Chang CC, Lerman OZ, et al. A murine model for studying diffusely injected human fat. Plast Reconstr Surg. 2009;124:74–81. 7. Chung MT, Hyun JS, Lo DD, et al. Micro-computed tomography evaluation of human fat grafts in nude mice. Tissue Eng Part C Methods 2013;19:227–232.
The Vasculosome Theory Sir: read with great interest the article entitled “True and ‘Choke’ Anastomoses between Perforator Angiosomes: Part II. Dynamic Thermographic Identification,” by Chubb et al. Interestingly, the findings of this study could better be explained by the “perforasome theory” rather than the “angiosome theory.”1 Both theories provide a guide to identify tissue units that can be reliably transferred in a single stage, based on a single
I
source of blood supply. However, both theories cannot reliably predict the exact quantity of tissue that can be supported by a single feeding vessel.2 Taylor and Palmer defined the angiosome as a vascular territory, which is marked by the presence of reduced caliber vessels in its periphery.3 The caliber by definition is narrow enough not to allow independent perfusion of the adjacent angiosome. Eventually, Taylor et al. also defined a perforasome as an inherent part of the angiosome.4 In the current article, Chubb et al. investigate reduced caliber vessels connecting perforasomes.5 It seems to contradict their angiosome theory, because by definition there cannot be narrow caliber vessels within an angiosome. The only reasonable explanation to sustain the description by Taylor et al. of a perforasome being a part of the angiosome, is to concede that angiosomes are not defined territories by themselves but are territories defined by several perforasomes (i.e., ∑perforasomes = an angiosome). Saint-Cyr et al. state, “Each perforator has its own vascular territory, called a perforasome, which carries a multidirectional flow pattern that is highly variable and complex.”1 Each perforasome is connected to its adjacent perforasomes by means of linking vessels, which could be “large-caliber” or “narrow-caliber” channels. However, the direction in which a larger tissue territory could be harvested based on a single perforator cannot be predicted based on the perforasome theory. McGregor and Morgan defined an axial flap as “a single pedicled flap which has an anatomically recognized arteriovenous system running along its long axis.”6 Branches of adjacent perforators, connected together by large-caliber channels, can form such a vascular axis. A unit of tissue incorporating this vascular axis and nourished by a single “feeder vessel” or perforator could be considered to be a “vasculosome.” In other words, any feeder vessel arising from a source vessel and contributing to the “vascular axis” should be sufficient to support the entire vasculosome independently. In fact, the objective of “flap delay” is to induce the formation of a neovasculosome with a centrally placed “vascular axis.”7 For a cutaneous vasculosome, suprafascial directionality of a perforator points toward the large-caliber channels with which it forms a vascular axis in the suprafascial plane.8 A single perforator supplying this vascular axis should be adequate to support a flap including the two (or more) adjacent “skin-perforasomes,”9 which will constitute a vasculosome (Fig. 1). Chubb et al. have suggested the use of thermography for detecting this vascular axis to aid flap elevation.5 Indeed, accurate mapping of a particular cutaneous vascular axis will result in better delineation of the cutaneous vasculosome and thus elevation of a more reliable flap. The muscle vasculosome has a vascular axis in the muscle. A single feeding vessel supplying any part of the muscle can nourish the entire muscle, provided that a vascular axis exists within the muscle (Fig. 2). If there are “narrow-caliber” channels linking adjacent muscle units, reliable harvest of both territories based on a single feeder is unlikely.10 Even so-called traditional minor pedicles can
449e
