Acta Astronautica 60 (2007) 383 – 390 www.elsevier.com/locate/actaastro

Can ultrasound counteract bone loss? Effect of low-intensity ultrasound stimulation on a model of osteoclastic precursor Monica Monicia,∗ , Pietro Antonio Bernabeib , Venere Basilec , Giovanni Romanoc , Antonio Contic , Luca Breschid , Leonardo Masottid , Augusto Cogolie a Consorzio CEO – Center of Excellence in Optronics, Largo Enrico Fermi 6, I-50125 Florence, Italy b Haematology Division, Careggi Hospital, V.le Morgagni 85, I-50134 Florence, Italy c Department of Clinical Physiopathology, University of Florence, V.le Pieraccini 6, I-50139 Florence, Italy d Department of Electronics and Telecommunications, University of Florence, Via S. Marta 3, I-50139 Florence, Italy e Space Biology Group, ETH-Technopark, Technoparkstr. 1, CH-8005 Zurich, Switzerland

Available online 23 October 2006

Abstract The aim of the present work is to determine whether mechanical stress caused by ultrasound (US) exposure affects osteoclastic precursor cells, thus addressing the hypothesis that mechanical strain-induced perturbation of preosteoclastic cell machinery can contribute to the occurrence of bone turnover alterations. Moreover, cell cytoskeleton was studied because of its supposed involvement in cell mechanotransduction. Our experimental model was the FLG 29.1 human cell line, previously characterized as an osteoclastic precursor model. Cell proliferation was quantified by trypan blue exclusion assay. Cell morpho-functional state was monitored by multispectral imaging autofluorescence microscopy. The expression of cytoskeletal components and markers of proliferation (Ki67) and osteoclastic differentiation (RANK) was analysed by immunocytochemistry. The findings demonstrated that US stimulation affects FLG 29.1 cell growth, depresses the expression of cytoskeletal components and markers of proliferation and differentiation, induces cell damage, thus supporting the hypothesis that US exposure inhibits osteoclastogenesis. These results have been compared with those obtained previously by exposure of FLG 29.1 cells to modelled hypogravity conditions. Finally, the possibility to utilize US stimulation for counteracting osteoporosis has been discussed. © 2006 Elsevier Ltd. All rights reserved. Keywords: Ultrasound; Osteoporosis; Hypogravity; Osteoclastic precursor; Osteoclastic differentiation

1. Introduction Since Galileo Galilei [1] realized that the rigidity of the skeleton is related to its load bearing function, the ∗ Corresponding author. Tel.: +39 55 427 1217;

fax: +39 55 427 1413. E-mail addresses: [email protected], monica.monici@unifi.it (M. Monici). 0094-5765/$ - see front matter © 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.actaastro.2006.09.023

influence of gravitational and mechanical stress on structure and homeostasis of bone has been widely studied [2,3]. Nevertheless, the molecular mechanisms involved at cellular level mostly remain to be understood. Experiments performed in altered gravitational conditions as well as the use of devices developed in order to expose the cells to different types of controlled mechanical stimuli [4] may help in studying mechanotransduction and consequent cell response. A great quantity

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of literature deals with the effect of gravitational and mechanical stress on osteoblasts and osteoclasts, the two cell populations responsible for bone turnover [5–7]. Our research concerns the behaviour of an osteoclastic precursor model (FLG 29.1 cell line) exposed to gravitational as well as mechanical stimuli. The final objective is to ascertain whether major genetic programs, as proliferation, differentiation and apoptosis, are directly affected in preosteoclastic cells, thus addressing the hypothesis that perturbation of preosteoclastic cell machinery by mechanical strains can play a role in bone turnover alteration. In previous studies, we found that FLG 29.1 cells cultured in hypogravity conditions, modelled by a random positioning machine, showed changes of cell morphology and metabolism, bone resorption activity, altered expression of cytoskeletal components and osteoclastic markers [8]. In this paper we describe the results obtained studying the response of FLG 29.1 cells to mechanical stress, applied by an ultrasound transducer. Ultrasound (US) is high-frequency acoustic radiation. The acoustical pressure waves can transmit mechanical energy to cells and tissues. US has been chosen, among various types of mechanical stimulations (stretch, shear stress, etc.), because of the widespread clinical application. Clinical trials reported enhanced healing rates of wounds, such as venous ulcers [9] and fractures [10–12]. Moreover, US has been reported to improve the efficacy of implant materials stimulating bone regeneration and bio-absorption [13,14]. US has been proposed as a countermeasure against osteoporosis [15]. Indeed, both in vivo [16,17] and in vitro [18] studies demonstrated that low intensity US promotes bone formation. Despite the proved advantages in clinical application of US treatment and extensive studies regarding the effect of low-intensity US on cells, the molecular mechanisms involved in the response to US stimulation are far from clear. Recent studies have reported that US promotes DNA synthesis in human osteoblasts [19], increases the expression of insulin-like growth factor I (IGF-I), osteocalcin (OC) and bone sialoprotein (BSP) in bone marrow derived stromal cells (ST2 cells) [20], stimulates the expression of c-fos, cyclooxygenase-2 (COX-2) and bone matrix proteins in bone-forming UMR-106 cells [21], alters cytokine release from osteoblasts [15]. Yang et al. found that, in osteoblastic cells exposed to US, the surface expression of 2 5 and 1 integrins increased and actin cytoskeleton reorganized. The same authors observed that US inhibited RANKL plus M-CSF-induced osteoclastic differentiation from bone marrow stromal cells [22].

Here, cell growth, cytoskeleton components and expression of proliferation and differentiation markers in osteoclastic precursors exposed to US stimulation have been investigated. The results obtained are consistent with the inhibition of osteoclastogenesis. 2. Materials and methods 2.1. Experimental model The FLG 29.1 human cell line was derived and stabilized from a culture of bone marrow cells, collected from a patient affected with acute monoblastic leukemia (FAB: M5a) and then characterized as a model of osteoclastic precursor [23]. The cells were cultured in 250 ml flasks in RPMI 1640 (Sigma-Aldrich, Italy), supplemented with 10% fetal calf serum (FCS) (SigmaAldrich, Italy), at 37 ◦ C and 5% CO2 . 2.2. US stimulation FLG 29.1 cells were resuspended in fresh culture medium (cell density 5.5 × 104 /ml). US stimulation was performed in a sterile air cabinet (Gelaire, BSB4A), utilizing a 10 MHz immersion transducer (Model V312, Panametrics, Inc., Waltham, MA) driven by an ultrasonic pulser/receiver (Model 5052, Panametrics, Inc., Waltham, MA), pulse duration 120 ns, pulse repetition frequency (PRF) 1, 100 Hz and 1 kHz, intensities, respectively, 0.1, 10, 100 mW/cm2 (Ispta, spatial peak temporal average intensity), negative peak acoustic pressure 250 kPa, exposure times 5, 15, 30 and 60 min. The transducer head was covered by a sterile cap and then immersed vertically into a culture chamber containing the cell suspension, which was stirred continuously and gently. Controls were treated in the same way, but with the US generator switched off. 2.3. Cell growth Daily cell counts were performed with a Neubauer chamber to assess growth kinetics. Viability was evaluated by trypan blue exclusion assay. 2.4. Multispectral imaging autofluorescence microscopy (MIAM) The term “autofluorescence” (AF) is used to distinguish the intrinsic fluorescence of cells and tissues from the fluorescence obtained by treating specimens with exogenous fluorescent markers, which bind to cell and tissue structures.

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When excited with radiation of suitable wavelength, some cell and tissue components behave as endogenous fluorophores: they pass to an excited state and then decay to the ground state with loss of energy, part of which consists in fluorescence emission. Cell AF is related to the cellular energy metabolism and the AF pattern reflects the organization of the intracellular structures, in accordance with the presence, among the principal endogenous fluorophores, of structural proteins and molecules involved in cell metabolism, for example nicotinic and flavinic coenzymes [24], whose emission is considered an indicator of the intracellular redox state [25]. Therefore, AF monitoring and analysis provide information on cell morphology and function. Cell AF was analysed using an inverted epifluorescence microscope (Nikon Eclipse TE 2000 E) equipped with an oil-immersion CF-UV fluor 100× objective (N.A. 1.3), under 365 nm excitation from a filtered (10 nm bandwidth interference filter, 365FS10–25, Andover Corp. Salem, NU, USA), high-pressure mercury lamp (HBO 100 W, Osram). The AF signal, transmitted through a dichroic mirror at 400 nm (DM400, Nikon), was detected by a Hires IV digital CCD camera (DTA, Italy) equipped with a Kodak KAF261E detector (20 m, 512 × 512 pixels). AF imaging was accomplished using a motorized filter wheel, containing up to eight different interference filters, placed in front of the CCD detector. This allowed for multispectral sequential acquisition in different emission bands. The choice of the filter combination was made on the basis of the main spectral bands determined by preliminary analysis of the AF spectra. Both the CCD camera and the filter wheel were controlled by a modified routine running under Visa software (DTA, Italy). AF images were directly digitized in the CCD controller with 16 bit dynamics and transferred to the storage computer on a digital interface. The size of the field detected by a 100× objective was about 69 × 69 m (spatial calibration of 0.13 m pixel−1 ), as determined by imaging 6 m fluorescent micro-spheres (Molecular Probes, Leiden, The Netherlands). 2.5. Immunocytochemistry Samples exposed to US stimulation and controls were concentrated (cell density 106 /ml) and then diluted 1:20 with CytoRich Blue Preservative Fluid. An Autocyte PREP System (Autocyte, Inc., Burlington, NC, USA) was utilized to prepare cell monolayers of 13 mm diameter, subsequently air-dried at room temperature for 2 h. The expression of F-actin, -tubulin,

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vimentin, RANK, Ki67 was assayed by immunocytochemical staining: cell monolayers were incubated with the specific anti-human antibodies, F-actin (clone NH3, Ab cam Limited, Cambridge, UK), -tubulin (clone DM1A, Neomarkers, Fremont, CA, USA), vimentin (clone V9, BioGenex, San Ramon, CA, USA), RANK (rabbit polyclonal, Santa Cruz Biotechnology, Santa Cruz, CA, USA), Ki67 (clone MIB-1 DakoCytomation, Denmark A/S, Glostrup, DK). Immunostaining was performed using a streptavidin-biotin peroxidase system kit (Lab Vision Corporation, Fremont, CA, USA). The slides were developed with 3, 3 -diaminobenzidine (DAB, BioGenex), and the nuclei were counterstained with Mayer’s hematoxylin (Merck KgaA, Darmstadt, Germany). A negative control was carried out by replacing the primary antibody with non-immune mouse serum. Sections of human osteosarcoma were used as the positive control. The monolayers were examined by light microscopy with a 100× immersion oil objective. At least 100 cells per slide were scored. 2.6. Statistics The data were expressed as mean ± SEM. The significance of the difference between the US treated group and the control was assessed by ANOVA test. The difference was considered significant when p < 0.05. 3. Results After US stimulation, each sample was analysed as follows: (a) cell growth was evaluated by trypan blue exclusion assay at 0, 24, 48 h after the treatment; (b) MIAM analysis was carried out immediately after the treatment, in order to evaluate cell morpho-functional state directly on viable samples; (c) part of the cells was prepared for immunocytochemical assessment of protein expression. In particular, we examined the expression of RANK, a key factor of osteoclastic differentiation [26,27]; Ki67, a nuclear antigen that is present throughout the active cell cycle (G1, S, G2 and M phases) but absent in resting cells (G0), and thus associated with cell proliferation [28]; major cytoskeleton components, such as -tubulin, vimentin, F-actin, because of the supposed implication of cytoskeleton in cell response to mechanical stimuli [29]. 3.1. Cell growth When the US stimulation of the cells was performed with the lowest PRF (1 Hz) and short exposure time

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1000 Control

US 15 min.

US 60 min.

103 Cells/ml

800 600 400 200 0 0

24

48

Time (h) Fig. 1. Effect of the time exposure to US stimulation (PRF 1 Hz) on cell growth. Significant differences between the two treatments and in comparison with control were observed at 48 h (p < 0.05).

1000

Control

US 100 Hz

103 Cells/ml

800 600 400 200 0 0

24

48

Time (h) Fig. 2. Effect of US stimulation (PRF 100 HZ, exposure time 15 min) on cell growth. A significant decrease was observed at 48 h (p < 0.01 as compared with control).

(< 15 min), we observed an increase in cell proliferation at both 24 and 48 h, after the treatment. For exposure time longer than 15 min, we monitored a slight increase of cell growth at 24 h, followed by a significant decrease at 48 h (Fig. 1). With PRF of 100 Hz and 1 kHz, we found a proliferation decrease at 24 h, which became much more important at 48 h (Fig. 2). The proliferation decrease depended on both PRF and exposure time: the longer the exposure time and the higher the PRF, the stronger the decrease. 3.2. AF analysis The AF pattern of the cells was analysed after the US stimulation and compared with the control. A 365 nm excitation wavelength was chosen in order to excite at the same time both NAD(P)H and flavins (oxidized form). In these experimental conditions, the intracellular autofluorescent sites co-localize with mitochondria and, to a lesser extent, with lysosomes [30], because the considered fluorophores are highly concentrated in these organelles. On the contrary, the nuclear content is

Fig. 3. Autofluorescence imaging of FLG 29.1 cells. Very different autofluorescence patterns characterize control cells (A) and cells exposed to US stimulation (exposure time 60 min, PRF 1 Hz) (B). See the text for description. Excitation wavelength 365 nm. For each sample, three 40 nm wide (full width at half maximum) spectral bands peaked at about 450, 550 and 650 nm (450 FS 40–25, 550 FS 40–25 and 650 FS 40–25, respectively, Andover, Salem, NH, USA) were used in order to sequentially acquire three fluorescence images in blue, green and red, with integration times of about 5 s. Monochrome images were then combined in a single multicolor image using the RGB technique. The multicolor images were obtained by the Image Combine Channels algorithm of Corel PHOTO-PAINT v 6.0 software (Corel Corporation, Ottawa, Canada) after the identification of the three greyscale images (acquired at 650, 550 and 450 nm, respectively) with the RGB components. Bar 5 m. For colour images see the electronic verison.

not excited and the nucleus appears dark. As expected, the AF imaging of control FLG 29.1 cells showed (Fig. 3A) a cytoplasmic blue–green AF, much more intense in the cell periphery, where the organelles were localized, while the nucleus appeared quite completely dark. When the cells were exposed for times shorter than 15 min to US stimulation with low PRF (1 Hz), no significant changes were observed in the AF pattern. Increasing the exposure time, we monitored alterations of the cell AF pattern, which revealed damage of cell structures (Fig. 3B). The principal feature was a

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Protein expression (a.u)

120%

Control

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US 30 min.

100% 80% 60% 40% 20% 0% RANK

Tub

Vim

F Act

Ki67

Fig. 5. Effect of US stimulation (exposure time 30 min, PRF 1 kHz) on the expression of the cytoskeleton components and markers of cell growth and differentiation. An important decrease of protein expression can be observed (p < 0.0001 as compared with control).

Protein expression [a.u]

120% 100% 80% 60% 40% 20% 0% Control Fig. 4. Autofluorescence imaging of FLG 29.1 cells. Monochrome images (spectral band width 400–700 nm) of control cells (A) and cells exposed to US stimulation (exposure time 30 min, PRF 1 kHz) (B), see the text for description. Excitation wavelength 365 nm. Bar 5 m.

homogeneously diffused blue AF, consistent with the alteration of membrane permeability that caused the efflux of NAD(P)H from mitochondria to the cytoplasm, followed by the influx of the coenzyme molecules into the nucleus and binding to the nuclear proteins. The increased cell dimensions suggested the occurrence of swelling. When a higher PRF was used for the treatments, irreversible damages to the cell structures appeared evident even at exposure times lower than 15 min. Numerous cells showing shrinkage, dissolution of intracellular compartments and highly fluorescent spots, due to diffusion of endogenous fluorophores and chromatin condensation, were observed (Fig. 4). 3.3. Immunocitochemistry US stimulation performed utilizing low PRF and short exposure time did not affect the expression of

US 15 min. US 30 min. US 60 min.

Fig. 6. Effect of the time exposure to US stimulation (PRF 100 Hz) on RANK expression in FLG 29.1 cells.

the cytoskeletal components considered (-tubulin, vimentin, F-actin) or that of RANK and Ki67, markers of osteoclastic differentiation and proliferation, respectively. Indeed, the immunocytochemical analysis did not evidence meaningful differences between treated and control cells. On the contrary, a decreased expression of all the proteins examined was monitored when the PRF was increased or the exposure time extended (Fig. 5): the longer the US exposure, the stronger the decrease of protein expression. This relationship, shown in Fig. 6 with reference to RANK, was ascertained for all the proteins assayed, but the magnitude of the decrease was very different. For example, after 30 min of US stimulation with PRF 1 kHz, -tubulin was less than 5%, in comparison with the control, while Ki67 was about 55% of the control. 4. Discussion The results of our study demonstrated that FLG29.1 cells are very sensitive to US stimulation and their response depends on treatment conditions, showing a relationship with PRF and time exposure.

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The many different and sometimes discordant reports in the literature suggest that the effect of US stimulation is cell-type dependent. For example, US promotes proliferation in osteoblasts [31,32], cartilage cells [33] and fibroblasts [34], while it inhibits endothelial cell growth [35]. In our experiments, the slight increase in preosteoclastic cell proliferation observed with mild treatments (PRF 1 Hz, exposure time lower than 15 min) was replaced by a significant decrease in cell growth when PRF and/or exposure times increased. This effect resulted dose-dependent, in agreement with what has been found by other authors on US-induced inhibition of endothelial cell proliferation [35]. Moreover, immunocytochemistry demonstrated a dose-dependent, highly significant decrease in the expression of Ki67 and RANK. Ki67 is considered an index of proliferation [28], thus the drop in its expression matched the decreased cell growth resulting from the trypan blue exclusion assay. RANK, a TNF-receptor family member, is present on the plasma membrane of osteoclastic precursors. The binding between the receptor and its ligand RANKL activates signal transduction pathways that ultimately lead to osteoclast differentiation and increased osteoclastic activity [26,27]. The drastic fall in RANK expression (5% of the control) that we found in preosteoclastic cells after US stimulation could be partially responsible for the US-caused inhibition of RANKL and M-CSF-induced osteoclastic differentiation observed by other authors [22]. Immunocytochemical staining also revealed a decrease in the expression of cytoskeletal proteins. -tubulin, vimentin and F-actin expression decreased to 4%, 25% and 40% of the controls, respectively. Cytoskeleton disorganization and/or reorganization is a recurrent feature in cells exposed to gravitational and mechanical stimuli, thus supporting the widely accepted hypothesis that the cytoskeleton is involved in cell gravisensing and mechanotransduction [36,29]. It is also known that important cytoskeletal alterations can lead to redistribution and damage of intracellular organelles and even to cell death [37]. AF analysis of FLG 29.1 cells, performed immediately after US exposure, was consistent with the occurrence of cell damage when a PRF higher than 1 Hz was utilized for the treatments, and these were protracted for more than 15 min. In particular, the AF pattern, showing a homogeneously distributed blue fluorescence, suggested the onset of membrane alterations and, consequently, mitochondrial damage. When the higher PRF (1 kHz) and exposure times longer than 30 min were applied, the presence of large, disorganized, intensely

fluorescent patches revealed the dissolution of intracellular compartments. In summary, apart from a slight increase in proliferation observed at PRF 1 Hz and short time exposure (< 15 min), which could be due to a synchronization of the cell cycle related to a functional recovery of injured cells, US stimulation inhibited preosteoclastic cell growth, decreased the expression of cytoskeletal components and markers of cell growth and differentiation, and induced cell damage. Such results fit very well into a framework of US-induced depression of osteoclastogenesis, as already described by other authors [22]. The inhibition of osteoclastogenesis could represent an important aspect in the improvement of fracture healing observed under US treatment. It is interesting to compare these findings with results previously obtained by exposure of FLG 29.1 cells to modelled hypogravity conditions, indicating an enhanced expression of major osteoclastic markers consistent with the triggering of a differentiation process [8]. The body of the literature indicates that unloading inhibits the osteoblastic function and stimulates the osteoclastic one; the contrary occurs when loading is applied. Thus unloading leads to bone loss, while loading promotes bone formation. US exposure is an easy and non-invasive means for applying mechanical stimulation in vivo. Indeed, in vivo studies and randomized, controlled clinical trials clearly proved a significant increase in the rate of bone healing and restoration of biomechanical function following US treatments [11]. Enhanced bone healing in tibial [10], radial [11], scaphoid [38] and mandibular [39] fractures has been observed. Studies carried out on animal models of ligament, tendon and cartilage injury demonstrated the beneficial effects of US stimulation in wound repair of tissues other than bone [40–42]. In US exposed repaired Achilles tendons, an increased order in the extracellular matrix network and collagen fibres distribution has been found [43]. Notwithstanding these positive indications, the application of US stimulation in vivo is limited, as the mechanisms involved in the induction of cellular response leading to the multifunctional effects described as well as the optimum exposure conditions are poorly understood. In the present study, we found a strong, dosedependent decrease in RANK expression in osteoclastic precursors exposed to US. The downregulation of RANK leads to an impairment in osteoclastic function, because the binding between the membrane receptor RANK and its ligand RANKL is a key passage in osteoclastic maturation. Recent in vitro studies showed that US stimulation decreases the osteoblastic

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secretion of cytokines involved in osteoclast differentiation and activation, thus depressing osteoclast function via paracrine regulation [15]. These findings suggest that US stimulation could inhibit bone resorption at least by two different mechanisms. This fact could be of consequence in the application of US not only for fracture healing but also for treating other bone diseases, such as osteoporosis. In conclusion, the possibility of inducing enhancement of the osteoblastic function and impairment of the osteoclastic one at the same time makes US a potential tool to counteract osteoporosis, providing that our knowledge on the molecular mechanisms involved in US-induced effects increases. In-depth research is needed to answer the question “Could US counteract bone loss?” and the study reported in this paper is only a very small contribution towards this goal. Acknowledgements The authors thank Prof. E. Biagi, Prof. F. Fusi and Dr. M. Paglierani for discussions and suggestions regarding their study. This work was supported by ASI—the Italian Space Agency. References [1] G. Galilei, Two new sciences, in: S. Drake, The Second Day, (Trans.), The University of Winsconsin Press, 1974, p. 109–146. [2] E.R. Morey-Holton, R.T. Wahlen, S.B. Arnaud, M. Van Der Meulen, The skeleton and its adaptation to gravity, in: M.J. Fregly, C.M. Blatteis (Eds.), Handbook of Physiology, vol. 1, Oxford University Press, Oxford, 1996, p. 691–719. [3] H.M. Frost, Vital biomechanics: proposed general concepts for skeletal adaptations to mechanical usage, Calcified Tissue International 42 (1988) 145–156. [4] T.D. Brown, Techniques for mechanical stimulation of cells in vitro: a review, Journal of Biomechanics 33 (2000) 3–14. [5] G. Carmeliet, L. Vico, R. Bouillon, Space flight: a challenge for normal bone homeostasis, Critical Reviews in Eukaryotic Gene Expression 11 (2001) 131–144. [6] J. Nagatomi, B.P. Arulandam, D.W. Metzger, A. Meuinier, R. Bizios, Frequency- and duration-dependent effects of cyclic pressure on select bone cell functions, Tissue Engineering 7 (2001) 717–728. [7] K. Kurata, T. Ucmura, A. Nemoto, T. Tateishi, T. Muratami, H. Higaki, H. Miura, Y. Iwamoto, Mechanical strain effect on bone-resorbing activity and messenger RNA expressions of marker enzymes in isolated osteoclast culture, Journal of Bone and Mineral Research 16 (2001) 722–730. [8] M. Monici, G. Agati, F. Fusi, M. Paglierani, A. Cogoli, P.A. Bernabei, Gravitational unloading induces osteoclast-like differentiation of FLG 29.1 cells, Journal of Gravitational Physiology 9 (2002) 261–262. [9] F. Johannsen, A.N. Gam, T. Karlsmark, Ultrasound therapy in chronic leg ulceration: a meta-analysis, Wound Repair and Regeneration 6 (1998) 121–126.

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