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Journal of Dermatological Science 69 (2013) 187–194

Association between collagen production and mechanical stretching in dermalextracellular matrix: In vivo effect of cross-linked hyaluronic acid filler.A randomised, placebo-controlled study

Virginie Turlier a,*, Alexandre Delalleau a, Christiane Casas a, Amandine Rouquier a, Pascale Bianchi a,Sandrine Alvarez a, Gwendal Josse a, Alain Briant b, Serge Dahan b, Christine Saint-Martory a,Jennifer Theunis a, Amel Bensafi-Benaouda a, Arnaud Degouy a, Anne-Marie Schmitt a, Daniel Redoules a

a Centre de Recherche sur la Peau Pierre Fabre, Hotel Dieu, Toulouse, Franceb Dermatologists, Toulouse, France

A R T I C L E I N F O

Article history:

Received 15 March 2012

Received in revised form 18 December 2012

Accepted 26 December 2012

Keywords:

Hyaluronic acid fillers

Extracellular matrix

Fibroblasts

Mechanical stretching

Neocollagenesis

A B S T R A C T

Background: The effects of hyaluronic acid (HA) injection on tissue collagen anabolism are suggested to

be related to the induction of mechanical stress, causing biochemical changes in skin physiology.

Objectives: To ascertain the association between dermal mechanics modulated by a hyaluronic acid-

based filler effect and metabolism.

Methods: Sixty females were randomised to receive a 0.5 mL injection of HA gel or isotonic sodium

chloride (control) in the arm. Skin biopsies were taken at baseline and after 1, 3 and 6 months. Protein

and gene expression of procollagen, matrix metalloproteinases (MMP) and MMP tissue inhibitors

(TIMP1) were measured blind by ELISA and qPCR, respectively. Injected volumes were measured by

high-frequency ultrasound and radiofrequency analysis. Skin layer effects of injections were analysed by

finite element digital modelling.

Results: One month after injection, the filler induced an increase in procollagen (p = 0.0016) and TIMP-1

(p = 0.0485) levels and relative gene expression of procollagen III and I isoforms compared with the

controls. After 3 months, procollagen levels remained greater than in the controls (p = 0.0005), whereas

procollagen expression and TIMP-1 and MMP content were no longer different. Forty-three percent of

the injected filler volume was found at 1 month, 26% after 3 months and 20% after 6 months.

Limitations: The ultrasound imaging technique limited the scope of the investigation and precluded an

evaluation of the action of the filler at the hypodermic level.

Conclusions: Integrating both mechanical and biological aspects, our results suggest that mechanical

stress generated by cross-linked HA plays a role in dermal cell biochemical response.

� 2013 Japanese Society for Investigative Dermatology. Published by Elsevier Ireland Ltd. All rights

reserved.

Contents lists available at SciVerse ScienceDirect

Journal of Dermatological Science

jou r nal h o mep ag e: w ww .e lsev ier . co m / jds

1. Introduction

The biomechanical performance of the skin depends essentiallyon the integrity of the dermal extracellular matrix. This matrix isarranged in a dense network of interlinked collagen (types I and III)and elastin fibres containing water-retaining proteoglycan andglycan macro-aggregates. With age, the collagen network undergoesre-organisation (density, orientation), resulting in a reduction incollagen and proteoglycan content [1], accompanied by an increasein denatured and degraded collagen [2]. These changes lead to a loss

* Corresponding author at: Centre de Recherche sur la Peau Pierre Fabre, Hotel Dieu,

2 rue Viguerie, BP3071, 31025 Toulouse cedex 3, France. Tel.: +33 5 62 48 85 13;

fax: +33 5 62 48 85 99.

E-mail address: [email protected] (V. Turlier).

0923-1811/$36.00 � 2013 Japanese Society for Investigative Dermatology. Published b

http://dx.doi.org/10.1016/j.jdermsci.2012.12.006

of matrix volume and, consequently, to a deterioration in mechani-cal properties [3], which may induce changes in cell metabolism.Indeed, a link between the mechanical stress exerted on differenttypes of tissues and neocollagen production has been suggested.Well-known examples are provided by studies on the adaptiveresponse of articular tissue which show that certain types of strainclearly stimulate the anabolic response [4]. Likewise, experimentaldata have confirmed the anabolic effect of mechanical strains on theextracellular matrix of fibroblasts cultured in a 3-dimensionalnetwork (collagen lattices) [5]. Furthermore, owing to the highdegree of biocompatibility of hyaluronic acid, the anabolic responsesof tissues after injection appear very different from those seen withother types of implants, which usually lead to the formation of afibrotic shell [6].

Hyaluronic acid (HA) fillers are reportedly effective in dermalcollagen anabolism, but data demonstrating efficacy are based

y Elsevier Ireland Ltd. All rights reserved.


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http://www.sciencedirect.com/science/journal/09231811


V. Turlier et al. / Journal of Dermatological Science 69 (2013) 187–194188

essentially on a single article [7]. In this study in 11 volunteers,Wang et al. demonstrated a significant increase in levels of dermalmRNA coding for collagen I, four and 13 weeks after injection of HAfiller, and in tissue inhibitors of metalloproteinases (TIMP)-1, -2and -3, four weeks after injection. The activating effect on collagen1 synthesis was confirmed by a histological approach. One of thebasic concepts advanced concerns the close relationship betweenstretching and biochemical changes in skin physiology. This stressappears to activate intracellular signalling pathways that modulatethe synthesis/degradation balance of the extracellular matrix [2].

The objective of this comparative placebo-controlled study wasto ascertain the association between metabolism and dermalmechanics modulated by a hyaluronic acid-based filler effect. Thechanges in the biochemical markers of fibroblast response wereanalysed over time and, in conjunction with this, we related thevolume of an injected filler, measured by 3D high-frequencyultrasound, to the mechanical stresses induced. This analysishighlights the strong links between the homeostasis of collagensynthesis and degradation pathways and the mechanical stressexerted by the injection of cross-linked HA.

2. Materials and methods

2.1. Study design and patient selection

This single-centre, comparative, randomised, placebo-con-trolled, in vivo, blind analysis study was conducted at the PierreFabre Skin Research Centre – CRP, Toulouse (France), in accordancewith the ethical principles of the Declaration of Helsinki and GoodClinical Practice guidelines. The protocol was approved by the Sud-Ouest et Outre Mer III Committee for the Protection of Persons(Ethics Committee) and the French Health Products Safety Agency(AFSSaPS). Each volunteer signed a written informed consent.

Sixty healthy female volunteers aged 35–75 years of skinphototype III with a body mass index in the normal range andhealthy arm skin were included.

Non-menopausal women must have been using effectivecontraception for at least 2 cycles before inclusion and have hada negative pregnancy test result at inclusion and at the end ofthe study. Menopausal women were not permitted to havemodified, introduced or stopped hormonal therapy in the 3months before inclusion or during the study. Subjects meetingthe following criteria were excluded: pregnant or breast-feedingwomen; history of allergy to cosmetics, hyaluronic acid or anyother ingredient of the test product, latex, or lidocaine- orprilocaine-medicated plaster; wound healing disorder; signs ofexcessive sun damage. Sun exposure within the month prior toinclusion or during the study, corticosteroids, retinoids, immu-nosuppressive or cardiovascular agents, or anticoagulantswithin the previous 3 months, and diuretics within the 2months preceding inclusion, were other reasons for non-inclusion, as were the use of topical corticosteroids or alpha-hydroxy acids at the test sites in the 4 months before inclusion,and the introduction or modification of hormonal treatmentsthat might affect the study results in the 3 months precedinginclusion or during the study.

2.2. Study products and injection technique

The study products were administered contralaterally in theposterior aspect of the arms by two trained investigators at theinjection visit (T0). The sides were randomly assigned in blocks of 4(using in-house software) by the department of Pre-industrialClinical Pharmacy, independently of our Skin Research Centre. Weused HA filler indicated for the treatment of moderate to deepwrinkles, such as nasolabial folds (NLFs).

The control was isotonic sodium chloride injection (0.9%) –Aguettant1.

A topical local anaesthetic cream (5% lidocaine and prilocaine)was applied to the injection sites 1 h before injection. The creamwas then removed and an injection of 0.5 ml of HA gel or controlwas administered using a linear threading technique over adistance of 2 cm (2� 0.25 ml).

2.3. Study period

The study was conducted over a 6-month period. Subjects weredivided into 3 groups of 20 (groups 1, 2 and 3), corresponding tothe time of the final biopsy: 1, 3 and 6 months post-injection,respectively. Allocation to the different groups was based onsubject availability and undertaken by the investigator during thepre-inclusion visit. The biopsy specimens (4 mm diameter) weretaken at T0 (pre-injection baseline value) and at the end of thestudy (Tfinal: 1, 3 or 6 months post-injection) from each injectedarea.

Ultrasound measurements were taken at each time point up toTfinal. The biopsies harvested for protein analysis were frozenimmediately at �80 8C and those for use for gene expressionanalysis were kept at 4 8C in RNAlater (Qiagen) overnight and thenstored at �20 8C until analysis.

2.4. Efficacy evaluation

The primary end-point was pro-collagen I expression 1, 3, 6months after injection, assayed by Elisa.

Secondary efficacy analysis included changes over time inTIMP1, MMP3 and MMP9 expression assayed by Elisa and geneexpression for collagen I (A1, A2), collagen III (A1), MMP3 andTIMP1 assayed by qPCR, volume and dispersion of the injectedhyaluronic acid calculated from ultrasound imaging, and defor-mation of the different skin layers evaluated by finite elementmodelling.

2.5. Protein expression of procollagen I, MMPs and TIMPs

To analyse the expression of procollagen I, MMP and TIMP, thebiopsy, dry-stored at �80 8C, was placed in PBS/0.1% Triton buffersolution and homogenised in the Tissue Lyser (Qiagen). Theresultant extract was centrifuged and the supernatant recovered.Procollagen I was assayed using the ELISA Procollagen Type I C-Peptide EIA kit (MK101, Takara) according to the supplier’srecommendations. TIMP-1 and MMP-3 and -9 were assayed inthe same original extract using the Fluorokine MAP Human TIMPpanel kit (LKT003, R&D) and the Fluorokine MAP MMP base kit(LMP000, R&D), respectively. The results obtained for each of thequantifications were standardised to the total protein concentra-tion determined in the biopsy extract by OPA assay (#26025,Thermo).

2.6. Gene expression

Total RNA was isolated using the iPrep PureLink kit (Invitrogen,Carlsbad, USA) according to the manufacturer’s directions,including DNase I digestion. After reverse transcription (10 mLof total RNA) with the High Capacity cDNA kit (Applied Biosystems,Foster City, USA), amplification reactions were performed on a7900 Real-time PCR System. The reaction volume per wellconsisted of 2.5 mL of cDNA diluted to one tenth, 5 mL of TaqManUniversal Master Mix (Life Technologies), 0.5 mL of 20 X Taqmanprobes (Gene expression assay, Life Technologies) and 2.5 mL ofRNase- and Dnase-free water. RPLP0 was used as endogenouscontrol. Results are expressed as DCt = 2(Ct RPLP0-Ct target gene).

V. Turlier et al. / Journal of Dermatological Science 69 (2013) 187–194 189

Primers and TaqMan probes used were designed by AppliedBiosystems (Life Technologies): COL1A1-Hs00164004_m1,COL1A2-Hs01028956_m1, COL3A1-Hs00943809_m1, MMP3-Hs00968308_m1 and TIMP1-Hs99999139_m1. Although they donot amplify genomic DNA (the assay probe spans an exon junction)a reverse transcription control without enzyme was stillperformed.

2.7. Ultrasound monitoring

High-frequency ultrasound at 20 MHz [8–10] enables thestructures of the dermis to be imaged by analysing the radio-frequency signal emitted in the skin by a transducer. The signalsreceived undergo morphological processing to map the echogeni-city of the area examined.

The apparatus used for this study was a Dermcup 2020(ATYS, France) coupled to a three-dimensional scanner tovisualise the volume of the region concerned. The acquisitionperformed, consisting of 300 2D images, allowed a volume of16 mm � 16 mm � 6.24 mm to be imaged at a resolution of550 � 300 � 268 voxels.

2.8. Calculation of volume and dispersion parameters

Manual contouring of the cavities containing the injectedhyaluronic acid was performed using a specifically developedsoftware platform. The injected volumes were depicted in threedimensions using VTK1 graphics libraries and by concatenatingthe contours generated using a simple linear interpolation of theexisting contours. The volume (V) and surface area (A) of theresultant object [11] were calculated and appeared essentialparameters for its characterisation. Dispersion of the object wascalculated by determining the ratio between the surface area (A) ofthe object and the theoretical area of a sphere of correspondingvolume (V). The sphere represents the theoretical minimum

Inclu ded a nd rand(february to octo

Group 1(1 mon th)N=20

Group 2 (3 m N=20

introd uction or change 2

Random isati on m

N=17

Absence of primary endpoint : 3

N=17

Per Protocol p

N=53

Fig. 1. Subject flow th

dispersion of the injected volume. The dispersion (D) could beexpressed as follows:

D ¼ A

ð36pV2Þ1=3(1)

This calculation allowed sensitive discrimination betweendifferent distributions of hyaluronic acid in the skin.

2.9. Finite element modelling

Finite element modelling [12] of this process was performed toanalyse the effects of an injection on the different skin layers. Inthis case, only the dermis and hypodermis, with respectivethicknesses of 1.22 mm (mean of each subject) and 2 mm, wereconsidered [13,14]. The system was reduced to two dimensions(plane strains). The geometry concerned corresponded to theexploratory field of the probe (16 mm). Meshing was obtained bysecond order triangular elements. To encourage lateral dispersion(generally observed on ultrasound), the injection was simulated byimposing excess hydrostatic pressure P = 0.1 kPa within anelliptical cavity with axes of 0.4 mm (diameter of injection needle)and 1 mm (chosen arbitrarily). Neo-Hookean hyperelastic behav-iour [15] was used to model the dermis and hypodermis, withrespective elastic constants of Xd

l ¼ 30 kPa and Xhl ¼ 10�2 kPa

[10,11]. A coefficient of compressibility of n = 0.45 was assumed,irrespective of the materials. The relationship between thisstructure and the underlying tissues was modelled throughrestrained boundary conditions of the nodes corresponding tothe inferior aspect of the hypodermis [13].

Because of the difference in viscosity of the two products, thedifferent sensation of volunteers during injection of the twoproducts, the appearance of the areas after injection (domed in thecase of HA) and the obvious differences between ultrasoundimages, the study cannot be blinded. However, to maintain some

omised : 60ber 20 09)

ont hs) Group 3 (6 months)N=20

therapy (HRT ) :

ist ake : 1 :

Premature w ithdr awal /Absence o f prima ry endpo int : 1

N=19

opulati on

rough the study.

0

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*** **

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TO Filler NaCI

Fig. 2. Expressions of protein markers and comparison of different time courses. (2a) Procollagen (2b) TIMP-1. Significance: *p < 0.05, **p < 0.01, ***p < 0.001. Procollagen

(2a) and TIMP-1 (2b) quantification by ELISA. The ELISA procedure is described in Section 2. Total protein concentration was used to normalise procollagen and quantify TIMP-

1. Procollagen was significantly increased 1 and 3 months after filler injection compared with the control (NaCl). One month after NaCl injection, the quantity of procollagen

was increased compared with baseline values. Filler injection increased TIMP-1 after one month compared with control and baseline values. Six months after NaCl injection,

TIMP-1 was increased compared with filler and baseline values.


integrity, biochemical analysis of biopsies were performed blind(label tubes hidden) by persons involved in this task only.

2.10. Statistical analysis

Statistical analysis was performed by a Biometry Unitindependent of the Pierre Fabre Skin Research Centre. Sixtysubjects, divided 20 to a group, were planned to be enrolled toensure a minimum sample size of 16 per group. The purpose of theanalysis was to compare treatments within each treatment groupand to analyse changes over time. The type I (a) risk was set at 0.05for all of the study. Analysis were performed on the per protocolpopulation (PP: randomised subjects receiving 1 dose of studymedication and not exhibiting any major protocol deviation).Statistical analysis was performed with SAS 8.2 software asfollows: the primary endpoint was evaluated by analysis ofvariance (paired t-test) for repeated measures on the changes frombaseline (T0), taking into consideration time, product, site,sequence and interactions as fixed effects and subject as a randomeffect factor. This analysis provided the results of intra- and inter-group comparisons. The same analysis was used for secondaryendpoints.

3. Results

3.1. Subjects

Sixty subjects were included from February to October 2009.One withdrew prematurely and six others exhibited a majorprotocol deviation (Fig. 1). Fifty-three subjects were analysed, withthe following numbers in each group: 1 month: 17, 3 months: 17,and 6 months: 19. The mean age was 53.5 � 13.8 years (median:44.0 years, minimum: 35; maximum: 73 years). Mean ages (�SD) forgroups 1, 2 and 3 were 55.35 (�13.12), 53.93 (�15.01) and 54.15(�14.22) years, respectively. There were no significant intergroupdifferences in subject characteristics and biochemical marker contentat baseline.

3.2. Protein markers

The values of the biochemical markers of the dermal matrixwere comparable in the 3 groups at inclusion. In group 1 (onemonth post-injection), a highly significant increase in procollagenlevels was found in the area treated with filler compared with the

control (+66%, p = 0.0016) and with baseline (+138%, p < 0.0001).Analysis showed that this increase was still present 3 months post-injection: +40% relative to the control side (p = 0.0005); +27%relative to the baseline content (p = 0.014) (Fig. 2a). Similarly,analysis of the changes from baseline at 1 month showed asignificant increase in TIMP-1 content in the filler group comparedwith the control (+19%, p = 0.0485) and with baseline (+28%,p = 0.0086) (Fig. 2b).

No difference was observed three months post-injection,whether between the treated and control areas or from baseline.Conversely, at 6 months, the opposite course was noted for TIMP-1content between treated and control areas, with a significantincrease for the control versus treated area (p = 0.032) and versusbaseline (p = 0.049) (Fig. 2b).

The mean dermal MMP-9 content (not presented here) did notdiffer significantly between treated and control areas in the 3groups. Similarly, no significant difference was apparent in theMMP-3 content measured in the treated and control areas onemonth after the injection. Conversely, the differences observedbetween these same two areas 3 months post-injection weresignificant (p = 0.044). Furthermore, inclusion of the MMP-3/TIMP-1 ratio revealed a statistically significant decrease between the twoproducts, expressed in the first month (p = 0.01) and still present at3 months (p = 0.022) (results not presented here).

3.3. Transcriptional markers

Quantitative RT-PCR provided additional evidence of the effectof an injection of cross-linked hyaluronic acid on the modulation ofprocollagen expression and substantiated the results obtainedusing the Elisa technique (Fig. 3). The results showed a significantincrease at 1 month in relative expression of the procollagen Iisoforms A1 and A2 (p = 0.007 and p = 0.0057, respectively) and ofprocollagen III (p = 0.004) at the sites treated by filler comparedwith those treated with control. This stimulant effect then declinedover time. Three months after injection of filler, a significantincrease from their baseline expression was still found in therelative expression of the A1 and A2 isoforms of procollagen I(p = 0.0266 and p = 0.0086, respectively) and procollagen III(p = 0.0126). By contrast, intergroup analysis failed to reveal asignificant difference from procollagen expression on the controlside.

An increase in relative TIMP1 expression at 1 month was alsofound at filler-treated sites compared with the control (p = 0.0135)

0

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T1 mon th T3 mon th T6 mon th

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a

** **

*

** **

**

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**

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b

c d

T0 Filler NaCl

Fig. 3. Expressions of transcriptional markers and comparison of different time courses. (3a) Col IA1 (3b) Col IA2 (3c) Col IIIA1 3d: TIMP-1. Significance: *p < 0.05, **p < 0.01,

***p < 0.001. Real-time reverse transcriptase-polymerase chain reaction (PCR) analysis of the genes encoding procollagen IA1 (3a), procollagen IA1 (3b), procollagen IIIA1 (3c)

and TIMP-1 (3d). Real-time PCR assay and isolation of total RNA are described in Section 2. mRNA levels were expressed normalised by the reference gene (RPLP0). Filler

injection produced an increase in procollagens at 1 and 3 months compared with control (NaCl). TIMP-1 relative expression was significantly increased 1 month after filler

injection compared with control and baseline values, and after NaCl injection compared with baseline. Six months after filler injection none of the markers expression

remained significantly increased compared with control.


and with baseline expression (p < 0.0001) (Fig. 3). However, asMMP-3 expression was below the limit of detection for half thesamples studied, these data were unevaluable.

3.4. Volume and dispersion

Only the ultrasound data for the patients from group 3 (6 monthfollow-up) were analysed to monitor the injections over time.Fifteen of these nineteen subjects were evaluable.

Fig. 4 shows a clear 2D illustration of an injected environment(a) (dark non-echogenic areas) versus a non-injected environ-ment (b). The majority of the images analysed showed the fillerto be localised in the hypodermis as far as its interface with thedermis (Fig. 4a). The mean individual variation was expressed asa percentage of the injected volume. Fig. 5 clearly demonstrates

Fig. 4. Example of 2D ultrasound images for (a) an injected and (b) a non-injected area. Th

the image. The injected HA appears as the non-echogenic (dark) area within the hypod

the changes due to the consumption and displacement ofhyaluronic acid over time. Following injection, a highly signifi-cant decrease (p < 0.0001) was observed over time (Figs. 5and 6a). Forty-three percent of the initial volume injected wasfound at 1 month and 26% and 20% after 3 and 6 months,respectively.

Concurrently with this decrease in volume, an increase wasnoted in the calculated dispersion values between the first andthird months (Figs. 5 and 6b). In qualitative terms, the volumeappeared less massive in terms of its organisation and the densityof the injected filler decreased. On the other hand, these dispersionvalues exhibited no significant change subsequently (Fig. 6b). Thissuggests that the appearance of the implant remained unchangedbetween 3 and 6 months, with hyaluronic acid simply beingdegraded according to the same distribution.

e dermis appears as the homogeneous, echogenic (bright) structure in the top half of

ermis.

Fig. 5. Comparison of volume dispersion at T1 month (a) and T3 months (b). Visualisation obtained using Paraview software. A loss in volume is shown, as well as an increase

in dispersion of the injection.


4. Discussion

This study was conducted to determine whether the mechani-cal stretching generated by HA injection could exert beneficialeffects on constituents of dermal extracellular matrix. 3Dultrasound enables the filler volumes to be visualised andquantified after injection. Owing to their identical structures, itwas not possible to distinguish strictly between injected andnative HA. Nevertheless, injected sites were clearly identified onthe images by the observation of hypo-echogenic areas.

Our analysis reveals that cross-linked HA accumulates in thehypodermis from its interface with the dermis, confirming anobservation made in the nasolabial folds [16]. The increase inpressure occasioned by the injection probably causes the filler toflow naturally towards an area that deforms more easily than thedermis, i.e. the hypodermis, which is much less rigid [13].

In addition, we had demonstrated by immunohistochemistry ina previous study that the filler tended to migrate towards the softerparts of the dermis [17]. We were not able to differentiate injectedHA gel strictly from native HA. Nevertheless, large hypo-echogenicareas were seen on high-frequency ultrasound images and largezones stained with Alcian blue were identified by histology. Theseobservations are characteristic of the presence of HA gel [18] andwere visible on the injected side only.

Finite element modelling was undertaken to corroborate thishypothesis (Fig. 7). This allows digital simulation of the action ofthe filler on the value and localisation of tissue stresses. Fig. 8shows clearly that, as a result of differences in the rigidity of the

0

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Vol

ume

(ml)

******

***

a

Fig. 6. Variations in mean injected volum

dermis and hypodermis, the material tends to flow towards thelatter structure. This appears to be the site of tensile stresseslocated predominantly in the superficial dermis. The changes inthese stresses occur quasi-linearly with the excess pressureapplied to the system. The values obtained and the deformationappear minor, but it is not possible to model the whole injectionbecause of the marked distortion of the meshing. The resultsobtained, correlated with the ultrasound measurements of volumeand dispersion of the filler, establish strong links betweenmechanical stresses and metabolic mechanisms.

Thus, the maximum volume and the minimum dispersion notedafter 1 month necessarily occasion greater mechanical stress in thesuperficial dermis. Similarly, while the dispersion remainsconstant, the decrease in volume between 3 and 6 months resultsin a reduction in these stresses. This effect is amplified by a long-term relaxation phenomenon described in the literature in relationto skin expanders for autologous grafts [19]. The reduction involume at 1, 3 and 6 months may be due to degradation of thehyaluronic acid injected or simply to migration, since theexploratory field of the probe precluded us from confirming onehypothesis or the other (limitation of the technique).

This stress induces deformation of the tissue matrix that mightaffect deformation-sensitive components on fibroblasts (mechan-oreceptors). In fact, mechanoreceptors convert the mechanicalsignal into an intracellular biochemical signal involving moleculartransformations that cause changes in the biochemical parameters.Transduction mechanisms have not been elucidated yet, butseveral articles suggest a role for integrins, in particular integrins

0

2

4

6

8


Dis

pers

ion

***b

e (a) and dispersion (b) over time.

Fig. 7. Meshing generated to simulate the effect of the injection on skin stresses (superficial dermis, papillary dermis and hypodermis, depicted in red, green and blue,

respectively). Injection was modelled by an elliptical area in which an increasing hydrostatic pressure load was applied.

Fig. 8. Finite element analysis (plane strain) of the effect of injection on (a) the vertical displacement field in deformed geometry (magnification 1�) and (b) tensile stress (s11

in MPa) in deformed geometry (magnification 10�).


a1b1 and a2b1, in the mechanical interaction between actin inthe cytoskeleton and collagen fibres in the perception ofmechanical stress [20,21].

The biochemical parameters sensitive to the mechanical effectare endogenous materials present in the dermis, predominantlycollagens, but also molecules that regulate the metabolism of thesematerials. The main catabolic actors are the enzymes involved inthe degradation of collagen, MMPs, and TIMPs, the latter beingmodulators of increased catabolic activity.

The results presented here show that the synthesis of neocolla-gen I and of procollagen I and III mRNA is significantly increased 1month after injection of the filler compared with baseline andcontrol. At the same time, a significant increase is observed in dermalTIMP-1 levels, confirmed by RT-PCR, and a significant decrease in theMMP-3/TIMP-1 ratio, raising the possibility of reduced proteolyticactivity in the dermis exposed to mechanical stress. However, aslight increase in procollagen I levels is observed on the control side 1month after injection compared with baseline. As with the filler, thissuggests a metabolic change induced by a modification of themechanical stress. However, it has been reported that the metabolicresponse to mechanical stress decreases with age and that this lackof response may be linked to the decrease in actin proteosynthesis,as has been established in isolated fibroblasts obtained from donorsof varying ages [22].

The ultrasound measurements at 3 months show a loss of fillervolume and an increase in its dispersion, testifying to a significantreduction in the stress exerted in the extracellular matrixmeasured 3 months after injection. At the same time, a markedreduction is noted in the metabolic response of the dermis,although the effect of the filler still persists at 3 months relative to

the baseline value and the control side. The increase induced inprocollagen I content by the filler is attenuated by comparison withthe variations observed in the first month, but the increasedexpression of the different procollagens remains significantrelative to the baseline values. In the case of TIMP1, a significantchange from baseline and from the control is no longer observed at3 months. On the other hand, with the MMP-3/TIMP-1 ratio, asignificant decrease is found at 3 months between filler-injectedsites, baseline and control.

At six months there is no longer any significant difference frombaseline except for TIMP-1. Similarly, ultrasound observations ofthe fate of the filler corroborate the existence of an increaseddispersion of the volume and validate the resultant loss ofcompression. This indicates a substantial reduction in themechanical stress exerted in the dermis. Thus, the changes inanabolic markers are accompanied by those in catabolic markers,the two processes being intimately linked.

To conclude, these results confirm those of Fisher and Voorhees[2] and further elucidate the effect of a mechanical action inducedby hyaluronic acid filling on dermal metabolism. Physiologically,the mechanical stresses can activate certain signal transductionpathways, causing the production of second messengers and othercell mediators. These events induce a change in the transcriptionalactivity of the fibroblast under stress and consequently of proteinmRNA levels in the extracellular matrix [5]. Another phenomenonassociated with the progressive degradation of hyaluronic acidmight contribute to a change in fibroblast metabolism. Thus,certain polysaccharide fragments can interact directly withfibroblasts via membrane receptors and in so doing engenderbeneficial anabolic effects [5].


Although the mechanism of action remains unclear, byintegrating both mechanical and biological aspects our studieshighlight the role that may be played by the mechanical stressgenerated by the filler on the biochemical response of dermalcells.

In fact, the age-related reduction in mechanical stretching dueto decreased HA content and fragmented collagen fibres in the skinis associated with a slowdown in biochemical response [1], so thatthe mechanical action of the filler would counteract this ageingphenomenon.

Role of the sponsor

The sponsor had no role in the design and conduct of the study,in the collection, analysis, and interpretation of data, or in thepreparation of the manuscript, review, or approval of themanuscript.

Financial disclosure

Drs. A. Briant, S. Dahan are private practitioners and served asconsultants and investigators for this study, for witch they receivedhonoraria. All other authors (V. Turlier, A. Delalleau, C. Casas,A. Rouquier, P. Bianchi, S. Alvarez, G. Josse, C. Saint-Martory,J. Theunis, A. Bensafi-Benaouda, A. Degouy, A.M. Schmitt,D. Redoules) are employed by Pierre Fabre Laboratories andreceived salaries, but they do not have any relevant financialinterests in the findings from this manuscript.

Funding

This study was supported by Pierre Fabre Dermocosmetique(PFDC).

Acknowledgements

We are grateful to Dr. Marielle Romet for providing editorialassistance on behalf of Laboratoires Pierre Fabre Dermo-Cosme-tique, and to Laboratoires Pierre Fabre Dermo-Cosmetique for theirfinancial support. We are grateful to Dr. Didier Coustou for help indesigning the study.

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