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Transplantation Surgery and Research

Understanding the Impact of Preservation Methods on the Integrity and Functionality of Placental Allografts

Johnson, Amy BS*; Gyurdieva, Alexandra MS*†; Dhall, Sandeep PhD*; Danilkovitch, Alla PhD*; Duan-Arnold, Yi PhD*‡

Author Information
doi: 10.1097/SAP.0000000000001101
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Human placental membranes (hPMs), most frequently amniotic membranes, have been used as biological dressings for wounds, burn treatment, and surgical reconstructive procedures since the early 20th century.1–3 The low immunogenicity and functional properties of hPMs make them an attractive choice for a biological wound dressing.4,5 Chronic wounds in particular benefit from the application of hPMs because of their anti-inflammatory, antioxidant, and angiogenic properties.6–11 In addition, hPMs are antifibrotic, antibacterial, and analgesic and help prevent wound desiccation, which are particularly important properties for treating burns.12,13 Human placental membranes have been successfully adapted to many reconstructive procedures because of their ability to conform to complex contoured surfaces and their inherent elasticity.14–17 Uses of hPMs include bladder, genital, and peritoneal reconstructive surgeries; oral surgeries; ocular surface reconstruction; reconstruction of dural defects; tendon, ligament, and synovial joint repairs; and prevention of surgical adhesions.13,15,18–20

These properties of hPMs are attributed to their diverse extracellular matrix (ECM), abundant bioactive factors, and endogenous neonatal cells, including epithelial cells, fibroblasts, and mesenchymal stem cells (MSCs).9,21–23 Methods of sterilization and preservation of fresh hPMs render them safe, off-the-shelf commercial products. However, these processes can lead to varying degrees of damage to the tissue components and therefore affect the functional properties of hPMs.24–26 Most commercial hPM products are devitalized (ie, contain dead cells that were killed during tissue preservation).27 Some studies have suggested that only preservation of placental matrix is important.28,29 However, it has been shown that viable amnion has a higher magnitude of anti-inflammatory, antioxidant, angiogenic, and chemoattractive effects as compared with devitalized amnion.6–8

The goal of this study was to compare the properties of hPMs processed by two preservation methods, cryopreservation and dehydration, which result in viable and devitalized tissue allograft products, respectively. The tissue integrity and functionality of commercial hPMs with one viable placental layer and with multiple devitalized layers were analyzed for differences in their composition, structural integrity, viability, ability to release growth factors, and dynamic response to in vitro hypoxia and inflammation.


Sample Preparation

Tissue Procurement and Ethics Statement

Human term placentas were provided by the National Disease Research Interchange (Philadelphia, Pa) and Cord Blood America, Inc (Las Vegas, Nev) from eligible donors after obtaining written, informed consent. Placental tissues were aseptically processed in a biological safety cabinet within 36 hours after collection and stored at 4°C until processing.

Aseptic Cryopreservation Process to Prepare Viable Cryopreserved Human Amniotic Membrane Samples

The amnion was separated from the umbilical cord and from the chorion by blunt dissection. Residual blood was removed from the amnion with Dulbecco phosphate-buffered saline (DPBS) (Thermo Fisher Scientific, Waltham, Mass), and the tissue was incubated in Dulbecco's modified Eagle medium (DMEM) (GE Healthcare Life Sciences, Piscataway, NJ) containing antibiotics at 37°C and 5% CO2 in a humidified atmosphere. Subsequently, antibiotics were removed by washing with DPBS, and the amnion was cut and mounted on a supportive backing.

Cryopreservation of the amnion was performed in a dimethyl sulfoxide (Mylan Inc, Canonsburg, Pa) containing cryoprotectant solution at a controlled cooling rate according to a proprietary process developed by Osiris Therapeutics, Inc. The tissue was then stored at −80°C and was thawed and rinsed with saline before use in the following experiments. This tissue will be referred to as vCHAM (viable cryopreserved human amniotic membrane) in this study, which is equivalent to commercially available Grafix Prime placental membrane allograft (Osiris Therapeutics, Inc, Columbia, Md).

Source of Dehydrated Human Amnion/Chorion Membrane

Dehydrated human amnion/chorion membrane (dHACM), commercially known as Epifix (MiMedix, Marietta, Ga), was used in this study. Samples were stored at room temperature and rehydrated with saline before use in the following experiments.

Comparison of Placental Tissue Components

Tissue Architecture and Composition Through Histological and Immunohistochemical Staining

Fresh human placental tissue (hPT), vCHAM, and dHACM samples were fixed in 10% formalin, embedded in paraffin, sectioned into 5-μm–thick cross-sections, and stained with hematoxylin and eosin according to standard protocols. The cytokeratin 18 and matrix metalloproteinase 9 (MMP9) content and tissue distribution in fresh hPT, vCHAM, and dHACM samples were determined by immunohistochemical staining using sheep primary antibodies (R&D Systems, Minneapolis, Minn) and antisheep secondary antibodies (Vector Laboratories, Burlingame, Calif) for cytokeratin 18 and goat primary antibodies (R&D Systems) and antigoat secondary antibodies (Vector Laboratories) for MMP9. Tissue sections incubated in the absence of primary antibodies served as a negative control. Histological and immunohistochemical staining and analysis were performed by an independent histology laboratory, Histoserv, Inc (Germantown, Md).

Cell Viability

An assessment of the cell viability of vCHAM and dHACM was performed using the LIVE/DEAD viability/cytotoxicity kit (Thermo Fisher Scientific) according to the manufacturer's instructions. Samples were then analyzed under a fluorescent microscope (Eclipse TE300; Nikon, Tokyo, Japan). Viable, intact cells within the tissue pieces were identified by green fluorescent calcein AM, whereas dead cells were labeled with red fluorescent ethidium homodimer-1. Green and red channel images were merged using ImageJ (National Institutes of Health, Bethesda, Md).

Characterization of Cells in Viable hPMs

Persistence of Viable Cells in a Chronic Wound Environment In Vitro

To evaluate the viability of cells within the tissue over time in vitro, vCHAM was incubated in low-glucose DMEM supplemented with 10% fetal bovine serum (FBS) (Thermo Fisher Scientific) at 37°C, 5% CO2, and 20% O2 in a humidified atmosphere for 4 and 7 days. To simulate an in vitro chronic wound environment, vCHAM was incubated in high-glucose DMEM supplemented with FBS, 10-ng/mL tumor necrosis factor α (TNF-α) (R&D Systems), and 100 ng/mL of bacterial lipopolysaccharide (LPS) (Sigma-Aldrich, St Louis, Mo) antigen at 37°C, 5% CO2, and 1% O2 in a humidified atmosphere for 4 and 7 days. The viability of vCHAM samples was then assessed using the LIVE/DEAD viability/cytotoxicity kit, and the samples were imaged and analyzed using fluorescent microscopy (Eclipse TE300). Three images of representative fields were taken of replicate wells at ×10 magnification. For each condition, the number of living cells and the number of dead cells were quantified by 3 blinded independent operators. The percent viability was calculated using the following formula: [number of viable cells / (number of viable cells + number of dead cells)]*100. The results of each operator were averaged.

Persistence of Viable Cells in an In Vivo Diabetic Mouse Chronic Wound

Animals (db/db mice purchased from Jackson Laboratories, Bar Harbor, Me) were housed at the Sobran BioScience vivarium at Johns Hopkins University. All experimental protocols were approved by the Sobran's Institutional Animal Care and Use Committee. The induction of chronic wounds in diabetic mice was performed as previously described.30 Briefly, 20 minutes before wounding, mice were treated once intraperitoneally with 3-amino-1,2,4-triazole (Sigma-Aldrich), an inhibitor of the antioxidant enzyme catalase, at 1-g/kg body weight. Mice were anesthetized using isofluorance, and a full-thickness 7-mm punch wound (excision of the skin and the underlying panniculus carnosus) was made on the midline of the back of the mouse after the hair had been removed with Nair (Church & Dwight Co, Inc, Ewing, NJ) 24 hours before wounding. The mouse wounds were treated once topically with mercaptosuccinic acid (Sigma-Aldrich), an inhibitor for glutathione peroxidase, at 150-mg/kg body weight. The wounds were then covered with Tegaderm (3 M, St Paul, Minn), a polyurethane dressing, for 20 days to prevent contamination. Mice with fully developed chronic wounds were used for the assessment of viable cell persistence after vCHAM application at day 20 post-wounding.

Before the application of vCHAM on in vivo chronic wounds, vCHAM was stained using VivoTrack 680 Imaging Agent (Perkin Elmer, Waltham, Mass) for 30 minutes at room temperature. Viable cryopreserved human amniotic membrane was applied on the chronic wounds and covered by Adaptic (Systagenix, Quincy, Mass), a nonadherent cellulose acetate dressing, followed by Tegaderm. Viable cryopreserved human amniotic membrane was removed at 4 and 8 days post-application and processed for cell isolation. Viable cryopreserved human amniotic membrane was washed with 30 mL of sterile DPBS 5 times and centrifuged at gradient speeds to remove wound fluid and exudate. Viable cryopreserved human amniotic membrane was then incubated in 0.125% trypsin (Thermo Fisher Scientific) for 10 minutes and chopped using a scalpel. Trypsin was inactivated by the addition of DMEM with 10% FBS. The digested tissue was strained through 40-μm cell strainer to collect cells. The isolated cells were imaged and analyzed using fluorescent microscopy (Eclipse TE300). Three images of representative fields were taken at ×10 magnification, and for each condition, the number of living cells was quantified. Freshly thawed vCHAM from the same donor was also digested and counted as described previously to provide a baseline preapplication viability. The percentage of viable cells that persisted after 4 and 8 days in the wound was calculated using the following formula: (number of viable cells post application / number of viable cells preapplication)*100.

Bromodeoxyuridine and Ki-67 Staining

To determine whether cells within fresh amnion and vCHAM tissue proliferate, the tissues were stained for bromodeoxyuridine (BrdU) and Ki-67. Fresh amnion and vCHAM were incubated in BrdU (Thermo Fisher Scientific) labeling solution (30-μg/mL BrdU in DMEM) for 24 hours at 37°C and 5% CO2 with humidity. The labeling solution was removed, and samples were washed with DPBS. The tissues were fixed with 4% paraformaldehyde in DPBS for 24 hours. Additional samples of fresh amnion and vCHAM were fixed in paraformaldehyde for BrdU staining without preincubation and for Ki-67 staining using rabbit primary antibodies (Abcam, Cambridge, Mass) and antirabbit secondary antibodies (Vector Laboratories). Hematoxylin was used to counterstain the nuclei. All samples were processed and stained by Histoserv, Inc.

Proliferation of Cells Isolated From vCHAM in Culture

Cells were released from equivalent-sized samples of fresh amnion and vCHAM by enzymatic digestion as previously described, quantified, seeded in Nunc tissue culture treated flasks (Thermo Fisher Scientific) in DMEM supplemented with 10% FBS, and incubated at 37°C and 5% CO2 with humidity.7 Representative images of the cellular confluence of the cultures were obtained after 14 days.

Functional Assays

Sustained Growth Factor Release

Equivalent-sized pieces of vCHAM and dHACM were incubated in low-serum culture medium (1% FBS in DMEM) for 3 hours (considered day 0) and 7 days at 37°C and 5% CO2 in a humidified atmosphere. Supernatants were collected, centrifuged, and stored at −80°C before testing. The samples were tested for platelet-derived growth factor BB (PDGF-BB), hepatocyte growth factor (HGF), and epidermal growth factor (EGF) by enzyme-linked immunosorbent assay (ELISA) (R&D Systems). Data are presented as mean (SD) for 1 representative experiment containing 3 replicates.

Effect of Chronic Wound Environment on Release of Angiogenic Vascular Endothelial Growth Factor In Vitro

Equivalent-sized pieces of vCHAM and dHACM were incubated in low-serum culture medium (1% FBS in DMEM) in a hypoxic environment (1% O2) with 10-ng/mL TNF-α and 100-ng/mL LPS for 96 hours at 37°C and 5% CO2 in a humidified atmosphere. Samples without TNF-α and LPS and incubated under normoxic (20% O2) conditions were considered baseline controls. Supernatants were collected, centrifuged, and then incubated with guanidine HCl (Thermo Fisher Scientific) containing calcium chloride (Sigma-Aldrich), Tris buffer (Sigma-Aldrich), and protease inhibitor cocktail (Roche, Basel, Switzerland) at 4°C overnight on a rotator. Samples were centrifuged and desalted using Zeba desalting columns (Thermo Fisher Scientific) per manufacturer's instructions. Desalted tissue lysates were collected and stored at −80°C until testing. The samples were tested for vascular endothelial growth factor (VEGF) by ELISA (R&D Systems). The guanidine HCl treatment of samples before measurement of VEGF was performed to eliminate factors that interfere with the detection of VEGF by ELISA. The relative percent change was calculated using the following formula: [(VEGF release in hypoxia − VEGF release at normoxia baseline control) / VEGF release at normoxia baseline control]*100. Data are presented as mean (SD) for 1 representative experiment containing 3 replicates. Student t test was used for statistical analysis, and P < 0.05 was considered significant.

Effect of Chronic Wound Environment on Release of Anti-inflammatory Prostaglandin E2 In Vitro

Equivalent-sized pieces of vCHAM and dHACM were incubated in low-serum culture medium (1% FBS in DMEM) in the absence or presence of 50 ng/mL of TNF-α for 20 hours at 37°C and 5% CO2 in a humidified atmosphere. Samples without the addition of TNF-α were considered baseline controls. Supernatant was collected, centrifuged, stored at −80°C, and thawed before testing. The levels of prostaglandin E2 (PGE2), an inhibitor of TNF-α, were quantified by ELISA (Cayman Chemical, Ann Arbor, Mich).31 The relative percent change was calculated using the following formula: [(PGE2 release in the presence of TNF-α − PGE2 release at baseline control) / PGE2 release at baseline control]*100. Data are presented as mean (SD) for 1 representative experiment containing 3 replicates. Student t test was used for statistical analysis, and P < 0.05 was considered significant.


Cryopreservation Maintains Native Tissue Structure

To determine the effect of hPM processing on tissue structure, vCHAM and dHACM were compared with fresh tissues using histological hematoxylin-and-eosin staining. Viable cryopreserved human amniotic membrane maintained tissue thickness and structural integrity comparable with the amniotic layer of fresh tissue (Figs. 1A, B). However, dehydration resulted in a tissue with significantly decreased matrix thickness and condensed matrix structure (Fig. 1C).

Hematoxylin and eosin staining of placental tissues. Histology of (A) fresh hPT displayed the fetal layers of the amnion, chorionic mesenchyme, and chorionic trophoblast and the choriodecidua where the fetal and maternal components of the placenta interface. B, vCHAM showed a thickness and structure similar to that of fresh hAM, whereas (C) dHACM showed a degradation of matrix and reduction in thickness. Images were taken at ×10 magnification for A and at ×40 magnification for B and C. hAM indicates amnion; hCD, choriodecidua; hCM, chorionic mesenchyme; hCT, chorionic trophoblast.

dHACM Contains Chorionic Trophoblast and Choriodecidual Layers

Immunohistochemical staining for cytokeratin 18, an epithelial cell intermediate filament that is highly expressed in trophoblast cells, confirmed that chorionic trophoblast and choriodecidua are present in dHACM.32 Fresh hPT has a layer of amniotic epithelial cells positive for cytokeratin 18 followed by negative amniotic and chorionic mesenchymal layers and a highly cytokeratin 18–positive chorionic trophoblast layer (Fig. 2A). Minimal positive staining for cytokeratin 18 was observed in the choriodecidua in comparison with the chorionic trophoblast. Equivalent to the amnion in fresh tissues, vCHAM was negative for cytokeratin 18 staining with the exception of the amniotic epithelial layer (Fig. 2B). However, the overall cytokeratin 18 staining pattern of dHACM was similar to that of fresh tissues confirming that dHACM composition includes the amnion, chorionic mesenchyme, chorionic trophoblast, and choriodecidua (Fig. 2C).

Cytokeratin 18 staining of placental tissues. A, In fresh hPT, the hAM epithelial layer and the hCT were positive for cytokeratin 18 (a trophoblast cell marker), and the hCD was weakly positive. The hAM mesenchymal layer and hCM were negative for cytokeratin 18; staining for cytokeratin 18 was similar in (B) vCHAM as fresh tissue with negative staining except in the epithelial cells. However, (C) the cytokeratin 18 staining pattern of dHACM was similar to that of the layers of fresh hPT with a distinct hCT layer. Images were taken at ×10 magnification for A and at ×40 magnification for B and C. hAM indicates amnion; hCD, choiodecidua; hCM, chorionic mesenchyme; hCT, chorionic trophoblast.

In addition to cytokeratin 18, we used MMP9 staining to confirm trophoblast and choriodecidua presence in dHACM because it has been shown that the trophoblast and choriodecidua in the placenta at term have high levels of MMP2 and MMP9, which are relevant to wound chronicity.33–35 Minimal levels of MMP9 were observed in the amniotic and chorionic mesenchymal layers of fresh hPT in comparison with the chorionic trophoblast and choriodecidua layers (Fig. 3A), which is in agreement with previously reported data.33 Viable cryopreserved human amniotic membrane, which contains only amnion, showed minimal MMP9 staining (Fig. 3B), whereas dHACM staining showed MMP9 distribution and content similar to that of fresh tissues (Fig. 3C).

MMP9 staining of placental tissues. In (A) fresh hPT, immunohistochemical staining displayed that the hAM and hCM contained a minimal content of MMP9 in comparison with the hCD and especially the hCT where MMP9 was mostly localized. B, vCHAM contained low MMP9 content in comparison with (C) dHACM that contained high MMP9 levels because of the presence of the hCT and hCD. Images were taken at ×10 magnification for A and at ×40 magnification for B and C. hAM indicates amnion; hCD, choriodecidua; hCM, chorionic mesenchyme; hCT, chorionic trophoblast.

Viable Cells Are Preserved in vCHAM Tissue

Placental membranes were tested for the presence of viable cells postthaw for vCHAM and after rehydration for dHACM using the LIVE/DEAD viability/cytotoxicity assay. Only vCHAM maintained a high degree of cellular viability similar to fresh amnion (Figs. 4A, B). Dehydrated human amnion/chorion membrane contained only dead cells (Fig. 4C).

Placental tissue viability and persistence of vCHAM viability in a chronic wound environment in vitro. Viable and dead cells in (A) fresh amnion, (B) just-thawed vCHAM, and (C) rehydrated dHACM were visualized microscopically after staining cells in the tissues using the LIVE/DEAD viability/cytotoxicity kit. Only (A) fresh amnion and (B) vCHAM contained viable cells. Viable and dead cells in vCHAM after 4 (D) and 7 (E) days in regular tissue culture conditions and after 4 (F) and (G) 7 days in a hypoxic, inflammatory, hyperglycemic environment were visualized microscopically after cell staining in the tissues using the LIVE/DEAD viability/cytotoxicity kit. Viable cryopreserved human amniotic membrane retained a high degree of viability for all conditions tested. Live cells stained green with calcein AM. Dead cells stained red with ethidium homodimer-1. All images were taken at ×10 magnification.

Viable cells in vCHAM persisted for 4- and 7-day periods when the membrane was thawed and placed in culture medium. Viable cryopreserved human amniotic membrane remained highly viable over time, with 91% viability after 4 days and 89% viability after 7 days (Figs. 4D, E). To mimic an in vitro chronic wound environment, vCHAM was placed in a hypoxic chamber with inflammatory TNF-α, bacterial LPS, and high-glucose medium. After 4 and 7 days of incubation, the cells in vCHAM exhibited an average of 76% and 60% viability, respectively (Figs. 4F, G).

Viable vCHAM Cells are Detectable in an In Vivo Diabetic Mouse Wound 4 and 8 Days After Application

The diabetic mouse wounds treated with inhibitors for antioxidant enzymes became chronic 3 weeks after wounding. Viable cryopreserved human amniotic membrane stained with VivoTrack 680 was applied onto the chronic wounds and then collected at days 4 and 8 after application. The isolated cells from these samples were positive for VivoTrack 680 staining (Figs. 5A, C). In addition, cells isolated from vCHAM seemed viable when stained with calcein AM (Figs. 5B, D), which demonstrates that vCHAM cells are capable of survival in diabetic wounds at least a week after application. The isolated Vivo Track prelabeled cells (the fluorescence of which was detected in the red channel) represent the total number of vCHAM cells collected after application. Of those, the cells positively stained for calcein AM represent viable vCHAM cells. The percentage of viable cells that persisted in the wound after 4 and 8 days compared with baseline (preapplication) was quantified as 38% and 8%, on average, respectively.

Viable cryopreserved human amniotic membrane cell persistence in an in vivo diabetic mouse model. Viable cryopreserved human amniotic membrane was prestained with VivoTrack 680 (a fluorescent dye that tracks cells in vivo) and placed on in vivo chronic wounds on db/db mice. After the collection of vCHAM from the wounds at days 4 or 8 after application, cells from vCHAM were isolated and stained with calcein AM (a fluorescent dye that detects viable cells). Stained cells were analyzed by fluorescent microscopy. Viable cryopreserved human amniotic membrane stained with (A, C) VivoTrack 680 showed cell persistence in the chronic wounds, whereas vCHAM stained with (B, D) calcein AM confirmed retention of viable cells at days 4 and 8 after vCHAM application to the wounds, respectively. All images were taken at ×10 magnification.

Cells in Cryopreserved Amnion Are Quiescent but Retain Proliferative Potential After Isolation From the Tissue

Neither fresh amnion nor vCHAM displayed any positive staining for BrdU or Ki-67 (Figs. 6A–D), as demonstrated by the absence of brown-stained cells in the samples. However, cells isolated from the tissue were successfully grown in a culture flask. After 14 days in culture, both fresh amnion and vCHAM grew to confluence and displayed comparable cell morphology (Figs. 6E, F).

Bromodeoxyuridine and Ki-67 stained amniotic membrane and proliferative capacity of amnion cells in culture. Fresh amnion (A, C) and vCHAM (B, D) were stained for BrdU and Ki-67, respectively, and then counterstained with hematoxylin. Neither tissue displayed positive staining (brown color) for either BrdU or Ki-67 indicating that both fresh amnion and vCHAM cells were quiescent within the tissue. Confluence of cells from (E) fresh amnion and (F) vCHAM after 14 days in tissue culture showed that they proliferated. Images A to D were taken at ×40 magnification, and images E and F were taken at ×10 magnification.

vCHAM Accumulation of PDGF-BB, HGF, and EGF Over Time

Viable cryopreserved human amniotic membrane and dHACM were incubated in culture medium, and the levels of multifunctional growth factors—PDGF-BB, HGF, and EGF—released in culture medium were measured for a period of 7 days. Viable cryopreserved human amniotic membrane, which has viable cells, had significantly higher levels of all 3 factors at day 7 compared with those at day 0 (baseline) (Figs. 7A–C). In vCHAM, PDGF-BB levels increased from 17 ± 3 to 304 ± 16 pg/mL (a 1721% increase), HGF levels increased from 17,418 ± 1447 to 141,206 ± 39,809 pg/mL (a 711% increase), and EGF levels increased from 144 ± 16 to 238 ± 32 pg/mL (a 65% increase). In dHACM, PDGF-BB levels increased from 32 ± 3 to 56 ± 18 pg/mL (a 77% increase), HGF levels increased from 33,128 ± 9264 to 76,692 ± 28,223 pg/mL (a 132% increase), and EGF levels decreased from 62 ± 12 to 58 ± 3 pg/mL (a 7% decrease).

The accumulation of growth factors after 7 days in culture for vCHAM versus dHACM. For (A) PDGF-BB, (B) HGF, and (C) EGF, vCHAM increased levels of growth factors released in culture medium over time to a greater extent than dHACM.

vCHAM Releases VEGF in Response to a Hypoxic, Inflammatory Environment In Vitro

The levels of the angiogenic factor VEGF in vCHAM and dHACM were measured after exposing the membranes to a hypoxic, inflammatory environment compared with baseline levels (normoxia, no inflammation). Only vCHAM responded with a 103 ± 34% increase in VEGF secretion upon stimulation (Fig. 8A). Dehydrated human amnion/chorion membrane displayed no response to stimulation with a change in VEGF levels of −11 ± 13%.

Placental tissue functional comparison. In comparison with dHACM, vCHAM showed functional superiority in (A) the increase in VEGF in response to hypoxia and inflammation and (B) the increase in PGE2 in response to inflammation.

vCHAM Releases PGE2 in Response to an Inflammatory Environment In Vitro

The anti-inflammatory activity of vCHAM and dHACM was tested by exposing the membranes to high levels of the proinflammatory cytokine TNF-α and measuring PGE2 released into the culture medium. Viable cryopreserved human amniotic membrane showed a significant increase in PGE2 from baseline upon TNF-α stimulation in comparison with dHACM (Fig. 8B). Viable cryopreserved human amniotic membrane increased the production of PGE2 by 350 ± 118%, whereas dHACM did not respond to TNF-α stimulation with a change in PGE2 levels of 8 ± 27%.


The objective of this study was to gain insight into how preservation methods affect placental membrane structure and function by comparing single-layer viable amnion, vCHAM, with multilayer nonviable dHACM. Differences in the composition and structural matrix integrity and in the functional properties between vCHAM and dHACM were investigated. Their functionalities were compared using assays that measured their anti-inflammatory and angiogenic activities and sustained growth factor release over time in vitro.

Viable cryopreserved human amniotic membrane is a cryopreserved hPM, which has been shown to be comparable with fresh tissue in structure and growth factor profile.25,27,28 Our results confirmed that cryopreserved amnion retains the structural integrity and thickness of fresh amnion. However, dHACM, which is a dehydrated tissue, showed alterations in matrix structure and a decrease in matrix thickness. These results confirm the findings previously reported in multiple studies. Both Rodríguez-Ares et al25 and Thomasen et al24 have reported decreased protein content in dehydrated amnion as compared with cryopreserved amnion. Thomasen et al24 detected only collagen IV and fibronectin in the basement membrane of dehydrated amnion, whereas cryopreserved amnion retained detectable levels of collagens IV to VII, fibronectin, and laminins indicating that ECM proteins are better protected via cryopreservation. Cooke et al27 showed that ECM structure was compromised in dehydrated amnion as compared with cryopreserved amnion, that dehydrated amnion contained lower levels of hyaluronic acid, and that dehydrated amnion lacked high–molecular-weight hyaluronic acid altogether. Overall, dehydration is a harsh preservation method that causes a loss of proteins, protein denaturation (leading to a decrease in protein functionality), and permanent damage to the structural and mechanical properties of amnion.25,27,36 Given that the ECM acts as a reservoir of growth factors and a regulator of cell function, dehydration can significantly decrease functionality of hPMs.37–39 In addition, irradiation, by which dHACM and many dehydrated allografts are sterilized, leads to a significant reduction in bioactive factor levels and damage to the ECM structure of hPMs.40

Commercial placental tissue allografts are regulated by the Federal Food and Drug Administration as human cell, tissue, and cellular and tissue-based products as defined in 21 CFR Part 1271 and Section 361 of the Public Health Services Act, which do not require premarket approval. Placental tissue allografts are categorized as a “skin substitute” and intended for homologous use as a wound cover to aid in the repair of acute and chronic wounds without restriction by etiology and locations. Therefore, placental allografts have a short time to market and a broad spectrum of clinical applications, which is an advantage in comparison with devices and drugs. Whereas most commercial placental membrane products available are composed solely of amnion, hPM products can contain different parts of the placenta—the amnion, chorion, umbilical cord, amniotic fluid, or a variety of their combinations—resulting in different product compositions and properties. Dehydrated human amnion/chorion membrane is marketed as being composed of both the amnion and chorion layers of the placenta.41 Human chorion has a mesenchymal or stromal layer containing neonatal fibroblasts and MSCs, growth factors, and ECM similar to that of the amnion stromal layer.21 As such, inclusion of the chorionic stromal layer can impart additional benefits to a biological wound product. However, chorion also includes the chorionic trophoblast layer, which contains high levels of MMPs and proinflammatory cytokines in the placenta at term.33,42,43 Xu et al33 have shown that the choriodecidua and trophoblast layers express MMP9, whereas amnion and chorion stromal layers express little to no MMP9, and amnion epithelium expresses some MMP9. We confirmed these findings by immunohistochemical staining of fresh hPT. Whereas the amnion and chorion stromal layers contained little to no evidence of MMP9 content, dHACM contained a layer of dense MMP9, further confirming that it contains trophoblast and choriodecidual layers. Thus, our cytokeratin 18 and MMP analysis established the presence of the trophoblast layer of chorion and choriodecidua in dHACM. Placental membranes are widely used for the treatment of chronic wounds, which have long been known to be characterized by an excess of MMPs.34,44–46 Recently, Gibson and Schultz46 have suggested that MMPs are also in excess in acute wound fluids and are related to impaired healing and dehiscence of surgically closed wounds. It has been shown that high levels of MMPs preclude wounds from healing by degrading growth factors and ECM proteins.46,47 One of the treatment strategies for chronic wounds includes the inhibition of MMPs.34,46 Therefore, the inclusion of trophoblast and choriodecidua rich in MMPs within placental products should be avoided.

Viable cryopreserved human amniotic membrane retained the structural and cellular integrity of fresh hPMs. We found that the cell viability of vCHAM postthaw was similar to that of fresh amnion and that dHACM contained only nonviable cells. This is consistent with previously reported results of cell viability of fresh amnion and viable cryopreserved amnion quantified using the trypan blue exclusion method—84% and 82% of viable cells were detected in fresh amnion and viable cryopreserved amnion, respectively.7 In addition, vCHAM retained viable cells in culture after 4 and 7 days in both standard tissue culture conditions and in an in vitro chronic wound environment simulated using hypoxia, inflammatory TNF-α, bacterial LPS, and high glucose. Furthermore, we confirmed viable cell persistence in vCHAM in vivo at days 4 and 8 after its application to diabetic mouse chronic wounds. This experiment was directed to address the fate of viable cells in vivo after vCHAM application to the wounds in mice. These data suggest that endogenous cells in vCHAM remain viable and can contribute to the biological activity of the graft. Our in vivo viable cell persistence data are in line with previously reported data.48,49 Wu et al49 found that 27% of culture-derived green fluorescent protein-expressing MSCs had persisted in a mouse impaired wound model at 7 days after application.

To further characterize the viable cells within vCHAM, BrdU and Ki-67 staining was performed and compared with fresh amnion. To our knowledge, the proliferative state of cells within fresh or cryopreserved amnion tissue has not been previously evaluated. Neither fresh amnion nor vCHAM showed BrdU- or Ki-67–positive staining, which indicates that the amnion cells are not proliferating within the tissue.50,51 However, when isolated and cultured, both fresh amnion and vCHAM proliferated and reached confluence in a tissue culture flask within 14 days. This is in agreement with previously reported results of fresh amnion cell isolation.52 Demonstration of the proliferative capacity and therefore the metabolic activity of cells isolated from cryopreserved amnion is a new finding of this study.

It is well documented that amniotic cells are capable of releasing a wide array of tissue reparative factors in vitro and in vivo.22,53–57 Thus, the presence of viable cells in vCHAM could serve as another source of growth factors in addition to growth factors present in the placental matrix. The release of 3 wound-relevant bioactive factors (PDGF-BB, HGF, and EGF) from vCHAM and dHACM was quantified. It has been shown that levels of growth factors such as PDGF and EGF are depleted in chronic ulcers, so a biological dressing that could provide PDGF and EGF to the wound bed should be beneficial.44,45,58,59 In comparison with dHACM, vCHAM increased growth factor levels after 7 days in culture demonstrating that the viable cells within vCHAM might be a source of bioactive factor accumulation.

The anti-inflammatory and angiogenic properties of amnion in vitro and in vivo have been well documented.1,7–9,11,55,60–62 Therefore, the release of anti-inflammatory and angiogenic factors by vCHAM and dHACM in response to TNF-α and hypoxia was investigated. A variety of growth factors in amnion contribute to its angiogenic activity such as basic fibroblast growth factor, PDGF, and VEGF.22,63 Duan-Arnold et al8 reported that the angiogenic activity of vCHAM in an endothelial cell tube formation assay in vitro decreased by approximately 73% when a VEGF neutralizing antibody was included in the assay. This indicates that VEGF is an important mediator of amnion angiogenic activity. In our study, we measured the angiogenic activity by VEGF release from vCHAM and dHACM upon exposure to hypoxia and inflammation to mimic a chronic wound environment. Exposure of dHACM to hypoxia did not increase VEGF levels compared with baseline (normoxia with no inflammation). In contrast to the lack of response by dHACM, exposure of vCHAM to hypoxia triggered an increase in VEGF. Multiple studies have confirmed that MSCs, one of the major cellular components of amnion, are able to up-regulate the release of VEGF when exposed to stressors such as TNF-α, LPS, and hypoxia.40,53,64,65 Thus, the observed increase in VEGF when vCHAM was exposed to hypoxic, inflammatory conditions was potentially due to the release of VEGF by amnion cells.

One mediator of the anti-inflammatory ability of vCHAM is the TNF-α inhibitor PGE2.31,55 Duan-Arnold et al7 previously reported that viable cryopreserved amnion increased PGE2 production upon TNF-α stimulation by 428% from baseline levels, whereas the PGE2 levels actually decreased in response to TNF-α in the nonviable cryopreserved amnion sample. Our study confirmed these previously reported results. We detected a 350% increase in PGE2 after exposure of vCHAM to proinflammatory TNF-α. However, when dHACM was exposed to TNF-α, only an 8% increase in PGE2 was observed, which is not statistically significant and is likely linked to variability in the amount of PGE2 in the matrix of different dHACM pieces. Given that amniotic cells such as epithelial cells and MSCs have been shown to be capable of releasing PGE2 in response to inflammation, our results suggest that the increase in PGE2 in the vCHAM cultures upon exposure to TNF-α could be mediated by viable cells.55

The results of this study confirm that cryopreservation is a gentler preservation method for hPMs than dehydration and indicate that retention of viable, active cells affects the functionality of hPMs in in vitro and in vivo wound models. Furthermore, we determined that the inclusion of multiple layers of hPMs does not seem to offset the degradation of the ECM and the lack of viable cells after tissue dehydration. This study demonstrates that using cryopreservation, a method in which all components of placental tissue are preserved including endogenous viable cells in their native state, results in better in vitro functionality of placental allografts. The clinical significance of viable skin substitutes remains to be investigated. However, recently, the first retrospective study comparing the effectiveness of vCHAM and dHACM for the management of acute or chronic wounds of various etiologies (venous leg ulcers, surgical wounds, diabetic foot ulcers, arterial ulcers, pressure ulcers, and other wounds) reported that, overall, 63.0% of wounds reached complete closure in the vCHAM group versus 18.2% in the dHACM group.66 Future prospective clinical trials comparing viable single-layer and nonviable multilayer allografts are required to confirm the results of this retrospective study.


The authors thank Dr Anthony Melchiorri of Osiris Therapeutics, Inc, for his valuable technical support and comments in reviewing this article.


1. Stern W. The grafting of preserved amniotic membrane to burned and ulcerated skin surfaces substituting skin grafts. JAMA. 1913;13:973–974.
2. Davis JW. Skin transplantation with a review of 550 cases at the Johns Hopkins Hospital. Johns Hopkins Med J. 1910;15:307.
3. Brindeau A. Création d'un vagin artificiel à l'aide des membranes ovulaires d'un oeuf à terme. Gynecol Obstet (Paris). 1934;29:385.
4. Akle CA, Adinolfi M, Welsh KI, et al. Immunogenicity of human amniotic epithelial cells after transplantation into volunteers. Lancet. 1981;2:1003–1005.
5. Kubo M, Sonoda Y, Muramatsu R, et al. Immunogenicity of human amniotic membrane in experimental xenotransplantation. Invest Ophthalmol Vis Sci. 2001;42:1539–1546.
6. Duan-Arnold Y, Gyurdieva A, Johnson A, et al. Soluble factors released by endogenous viable cells enhance the antioxidant and chemoattractive activities of cryopreserved amniotic membrane. Adv Wound Care (New Rochelle). 2015;4:329–338.
7. Duan-Arnold Y, Gyurdieva A, Johnson A, et al. Retention of endogenous viable cells enhances the anti-inflammatory activity of cryopreserved amnion. Adv Wound Care (New Rochelle). 2015;4:523–533.
8. Duan-Arnold Y, Uveges TE, Gyurdieva A, et al. Angiogenic potential of cryopreserved amniotic membrane is enhanced through retention of all tissue components in their native state. Adv Wound Care (New Rochelle). 2015;4:513–522.
9. Niknejad H, Peirovi H, Jorjani M, et al. Properties of the amniotic membrane for potential use in tissue engineering. Eur Cell Mater. 2008;15:88–99.
10. Lockington D, Agarwal P, Young D, et al. Antioxidant properties of amniotic membrane: novel observations from a pilot study. Can J Ophthalmol. 2014;49:426–430.
11. Faulk WP, Matthews R, Stevens PJ, et al. Human amnion as an adjunct in wound healing. Lancet. 1980;1:1156–1158.
12. Bose B. Burn wound dressing with human amniotic membrane. Ann R Coll Surg Engl. 1979;61:444–447.
13. Munoyath SK, Prasad K. Efficacy of human amniotic membrane and collagen in maxillofacial soft tissue defects—a comparative clinical study. J Oral Maxillofac Surg Med Pathol. 2015;27:786–790.
14. Mamede AC, Carvalho MJ, Abrantes AM, et al. Amniotic membrane: from structure and functions to clinical applications. Cell Tissue Res. 2012;349:447–458.
15. Fairbairn NG, Randolph MA, Redmond RW. The clinical applications of human amnion in plastic surgery. J Plast Reconstr Aesthet Surg. 2014;67:662–675.
16. Pennati G. Biomechanical properties of the human umbilical cord. Biorheology. 2001;38:355–366.
17. Oxlund H, Helmig R, Halaburt JT, et al. Biomechanical analysis of human chorioamniotic membranes. Eur J Obstet Gynecol Reprod Biol. 1990;34:247–255.
18. Lawson VG. Oral cavity reconstruction using pectoralis major muscle and amnion. Arch Otolaryngol. 1985;111:230–233.
19. Bleggi-Torres LF, Werner B, Piazza MJ. Ultrastructural study of the neovagina following the utilization of human amniotic membrane for treatment of congenital absence of the vagina. Braz J Med Biol Res. 1997;30:861–864.
20. Trelford JD, Trelford-Sauder M. The amnion in surgery, past and present. Am J Obstet Gynecol. 1979;134:833–845.
21. Parolini O, Alviano F, Bagnara GP, et al. Concise review: isolation and characterization of cells from human term placenta: outcome of the first international workshop on placenta derived stem cells. Stem Cells. 2008;26:300–311.
22. Maxson S, Lopez EA, Yoo D, et al. Concise review: role of mesenchymal stem cells in wound repair. Stem Cells Transl Med. 2012;1:142–149.
23. Hao Y, Ma DH, Hwang DG, et al. Identification of antiangiogenic and antiinflammatory proteins in human amniotic membrane. Cornea. 2000;19:348–352.
24. Thomasen H, Pauklin M, Steuhl KP, et al. Comparison of cryopreserved and air-dried human amniotic membrane for ophthalmologic applications. Graefes Arch Clin Exp Ophthalmol. 2009;247:1691–1700.
25. Rodríguez-Ares MT, López-Valladares MJ, Touriño R, et al. Effects of lyophilization on human amniotic membrane. Acta Ophthalmol. 2009;87:396–403.
26. von Versen-Hoeynck F, Steinfeld AP, Becker J, et al. Sterilization and preservation influence the biophysical properties of human amnion grafts. Biologicals. 2008;36(4):248–255.
27. Cooke M, Tan EK, Mandrycky C, et al. Comparison of cryopreserved amniotic membrane and umbilical cord tissue with dehydrated amniotic membrane/chorion tissue. J Wound Care. 2014;23:465–474.
28. Kruse FE, Joussen AM, Rohrschneider K, et al. Cryopreserved human amniotic membrane for ocular surface reconstruction. Graefes Arch Clin Exp Ophthalmol. 2000;238:68–75.
29. Ilic D, Vicovac L, Nikolic M, et al. Human amniotic membrane grafts in therapy of chronic non-healing wounds. Br Med Bull. 2016;117:59–67.
30. Dhall S, Do DC, Garcia M, et al. Generating and reversing chronic wounds in diabetic mice by manipulating wound redox parameters. J Diabetes Res. 2014;2014:562625.
31. Harris SG, Padilla J, Koumas L, et al. Prostaglandins as modulators of immunity. Trends Immunol. 2002;23:144–150.
32. Graham CH, Lysiak JJ, McCrae KR, et al. Localization of transforming growth factor-beta at the human fetal-maternal interface: role in trophoblast growth and differentiation. Biol Reprod. 1992;46:561–572.
33. Xu P, Alfaidy N, Challis JR. Expression of matrix metalloproteinase (MMP)-2 and MMP-9 in human placenta and fetal membranes in relation to preterm and term labor. J Clin Endocrinol Metab. 2002;87:1353–1361.
34. Amălinei C, Căruntu ID, Giuşcă SE, et al. Matrix metalloproteinases involvement in pathologic conditions. Rom J Morphol Embryol. 2010;51:215–228.
35. Armstrong DG, Jude EB. The role of matrix metalloproteinases in wound healing. J Am Podiatr Med Assoc. 2002;92:12–18.
36. Jiang S, Nail SL. Effect of process conditions on recovery of protein activity after freezing and freeze-drying. Eur J Pharm Biopharm. 1998;45:249–257.
37. Badylak SF. Regenerative medicine and developmental biology: the role of the extracellular matrix. Anat Rec B New Anat. 2005;287:36–41.
38. Agren MS, Werthén M. The extracellular matrix in wound healing: a closer look at therapeutics for chronic wounds. Int J Low Extrem Wounds. 2007;6:82–97.
39. Olczyk P, Mencner Ł, Komosinska-Vassev K. The role of the extracellular matrix components in cutaneous wound healing. Biomed Res Int. 2014;2014:747584.
40. Paolin A, Trojan D, Leonardi A, et al. Cytokine expression and ultrastructural alterations in fresh-frozen, freeze-dried and γ-irradiated human amniotic membranes. Cell Tissue Bank. 2016;17:399–406.
41. Available at: Accessed March 22, 2017.
42. Paradowska E, Blach-Olszewska Z, Gejdel E. Constitutive and induced cytokine production by human placenta and amniotic membrane at term. Placenta. 1997;18:441–446.
43. Steinborn A, Günes H, Halberstadt E. Signal for term parturition is of trophoblast and therefore of fetal origin. Prostaglandins. 1995;50:237–252.
44. Tarnuzzer RW, Schultz GS. Biochemical analysis of acute and chronic wound environments. Wound Repair Regen. 1996;4:321–325.
45. Trengove NJ, Stacey MC, MacAuley S, et al. Analysis of the acute and chronic wound environments: the role of proteases and their inhibitors. Wound Repair Regen. 1999;7:442–452.
46. Gibson DJ, Schultz GS. Molecular wound assessments: matrix metalloproteinases. Adv Wound Care (New Rochelle). 2013;2:18–23.
47. Martel-Pelletier J, McCollum R, Fujimoto N, et al. Excess of metalloproteases over tissue inhibitor of metalloprotease may contribute to cartilage degradation in osteoarthritis and rheumatoid arthritis. Lab Invest. 1994;70:807–815.
48. Kim SW, Zhang HZ, Guo L, et al. Amniotic mesenchymal stem cells enhance wound healing in diabetic NOD/SCID mice through high angiogenic and engraftment capabilities. PLoS One. 2012;7:e41105.
49. Wu Y, Chen L, Scott PG, et al. Mesenchymal stem cells enhance wound healing through differentiation and angiogenesis. Stem Cells. 2007;25:2648–2659.
50. van Diest PJ, Brugal G, Baak JP. Proliferation markers in tumours: interpretation and clinical value. J Clin Pathol. 1998;51:716–724.
51. Ohta Y, Ichimura K. Proliferation markers, proliferating cell nuclear antigen, Ki67, 5-bromo-2'-deoxyuridine, and cyclin D1 in mouse olfactory epithelium. Ann Otol Rhinol Laryngol. 2000;109:1046–1048.
52. Mihu CM, Rus Ciucă D, Soritău O, et al. Isolation and characterization of mesenchymal stem cells from the amniotic membrane. Rom J Morphol Embryol. 2009;50:73–77.
53. Crisostomo PR, Wang Y, Markel TA, et al. Human mesenchymal stem cells stimulated by TNF-alpha, LPS, or hypoxia produce growth factors by an NF kappa B- but not JNK-dependent mechanism. Am J Physiol Cell Physiol. 2008;294:C675–C682.
54. Kim SW, Zhang HZ, Kim CE, et al. Amniotic mesenchymal stem cells have robust angiogenic properties and are effective in treating hindlimb ischaemia. Cardiovasc Res. 2012;93:525–534.
55. Silini A, Parolini O, Huppertz B, et al. Soluble factors of amnion-derived cells in treatment of inflammatory and fibrotic pathologies. Curr Stem Cell Res Ther. 2013;8:6–14.
56. Payne WG, Wachtel TL, Smith CA, et al. Effect of amnion-derived cellular cytokine solution on healing of experimental partial-thickness burns. World J Surg. 2010;34:1663–1668.
57. Uberti MG, Ko F, Pierpont YN, et al. The use of amnion-derived cellular cytokine solution (ACCS) in accelerating closure of interstices in explanted meshed human skin grafts. Eplasty. 2009;9:e12.
58. Pierce GF, Tarpley JE, Tseng J, et al. Detection of platelet-derived growth factor (PDGF)-AA in actively healing human wounds treated with recombinant PDGF-BB and absence of PDGF in chronic nonhealing wounds. J Clin Invest. 1995;96:1336–1350.
59. Chen SM, Ward SI, Olutoye OO, et al. Ability of chronic wound fluids to degrade peptide growth factors is associated with increased levels of elastase activity and diminished levels of proteinase inhibitors. Wound Repair Regen. 1997;5:23–32.
60. Yamahara K, Harada K, Ohshima M, et al. Comparison of angiogenic, cytoprotective, and immunosuppressive properties of human amnion- and chorion-derived mesenchymal stem cells. PLoS One. 2014;9:e88319.
61. Egan TJ, O'Driscoll J, Thakar DR. Human amnion in the management of chronic ulceration of the lower limb: a clinico-pathologic study. Angiology. 1983;34:197–203.
62. Gruss JS, Jirsch DW. Human amniotic membrane: a versatile wound dressing. Can Med Assoc J. 1978;118:1237–1246.
63. Cui T, Kirsner R, Li J. Angiogenesis in chronic wounds. Adv Wound Care. 2010;1:347–352.
64. Lee EY, Xia Y, Kim WS, et al. Hypoxia-enhanced wound-healing function of adipose-derived stem cells: increase in stem cell proliferation and up-regulation of VEGF and bFGF. Wound Repair Regen. 2009;17:540–547.
65. Madrigal M, Rao KS, Riordan NH. A review of therapeutic effects of mesenchymal stem cell secretions and induction of secretory modification by different culture methods. J Transl Med. 2014;12:260.
66. Johnson EL, Marshall JT, Michael GM. A comparative outcomes analysis evaluating clinical effectiveness in two different human placental membrane products for wound management. Wound Repair Regen. 2016;25:145–149.

allografts; wounds; burns; reconstructive surgery; placental membranes; amnion; cryopreservation; dehydration; cell viability

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