Journal of Molecular Histology

, Volume 42, Issue 1, pp 47–58

Keratinocyte growth factor 1, fibroblast growth factor 2 and 10 in the healing tympanic membrane following perforation in rats


    • Ear Sciences Centre, School of SurgeryThe University of Western Australia
    • Ear Science Institute Australia
    • Department of ENT, E Block OutpatientsSir Charles Gairdner Hospital
  • Sharon L. Redmond
    • Ear Sciences Centre, School of SurgeryThe University of Western Australia
    • Ear Science Institute Australia
  • Marcus D. Atlas
    • Ear Sciences Centre, School of SurgeryThe University of Western Australia
    • Ear Science Institute Australia
  • Reza Ghassemifar
    • Department of Haematology, Experimental Haematology Research UnitPathWest Laboratory Medicine
    • The School of Pathology and Laboratory MedicineThe University of Western Australia
Original Paper

DOI: 10.1007/s10735-010-9306-2

Cite this article as:
Santa Maria, P.L., Redmond, S.L., Atlas, M.D. et al. J Mol Hist (2011) 42: 47. doi:10.1007/s10735-010-9306-2


The aim of this study was to provide a transcriptome profile of Keratinocyte Growth Factor (KGF)-1, Fibroblast Growth Factor (FGF) 2 and FGF10 (KGF2) in the healing rat tympanic membrane (TM) over 7 days and an immunohistochemical account over 14 days following perforation. KGF1, FGF2, and FGF10 play important roles in TM wound healing. The tympanic membranes of rats were perforated and sacrificed at time points over a 14-day period following perforation. The normalized signal intensities and immunohistochemical protein expression patterns at each time point for KGF1, FGF2, and FGF10 are presented. The primary role of both KGF1 and FGF2 appeared to be in the proliferation and migration of keratinocytes. Whereas the role of KGF1 appeared to be exclusively concerned with increased proliferation and migration at the perforation site, the continued expression of FGF2, beyond perforation closure, suggested it has an additional role to play. FGF10 (KGF2), whilst possessing the highest sequence homologous to KGF1, has a different role in TM wound healing. The effect of FGF10 on keratinocytes in wound healing appeared to emanate from the connective tissue layer.


KGFFGFKeratinocyte Growth FactorFibroblast Growth FactorTympanic MembraneRatWound healingHistologyMicroarray


The process of wound healing is complex with many growth factors involved. Some growth factors, including Fibroblast Growth Factors (FGFs) are believed to play critical roles in this process. FGFs are a family of 22 growth factors. They interact with heparin or heparin sulfate proteoglycans for added stability (Werner and Grose 2003). FGFs stimulate proliferation and differentiation of a variety of cell types including keratinocytes, fibroblasts, chondrocytes, adrenocortical cells, vascular smooth muscle, vascular endothelium, and neurons. FGFs stimulate cell proliferation by shortening the G1 phase of the cell cycle (Clymer et al. 1996). They also induce angiogenesis (Ma et al. 2002) and increase protease production leading to matrix remodelling (Clymer et al. 1996). It is important to note that growth factors often act in combination with little or no effect without another growth factor (Lynch et al. 1987). Only three of the nine FGFs were once thought to play a role in wound healing (Hom 1995). These are FGF1 (acidic FGF), FGF2 (basic FGF) and KGF1 (FGF7). However, previously published data reveals a role for FGF10 (also known as KGF2) similar to that of KGF1 (Braun et al. 2004).

There are several studies examining the role of FGF2 application to the acutely healing tympanic membrane (TM) (Mondain and Ryan 1994; Fina et al. 1991; Fina et al. 1993; Mondain et al. 1991; Vrabec et al. 1994). Research suggests that only high doses have a significant effect but these effects were counterbalanced by the effects of a hypertrophic wound, possibly due to the gelfoam™ used for application (Mondain et al. 1991). No significant differences were noted when applying a neutralising antibody to FGF2 to a healing TM (Mondain et al. 1995). Of the FGF family, only the immunohistochemistry of FGF2 has been previously examined in the healing TM (Mondain and Ryan 1995). FGF2 and KGF1 mRNA expression levels have been examined following TM perforation (Ishibashi et al. 1998). FGF2 was found to have no significantly different levels in human TM chronic perforations when examined via immunohistochemistry (Somers et al. 1998). The phenotypic expression of KGF1, FGF2 and FGF10 in cultured human TM explants has also been decribed (Redmond SL et al. 2010). When applied to healing TMs KGF1 is thought not to influence the rate of healing but to lead to a more organised wound repair process (Clymer et al. 1996). Whilst there are a number of attempts to use FGFs to influence TM wound healing, there is a lacking in the literature examining the roles of these factors in the normal healing process of the TM.

A detailed histological account of TM wound healing has been recently published (Santa Maria et al. 2010a). The aim of this study was to provide a transcriptome profile of KGF1, FGF2, and FGF10 in the healing TM in rats over 7 days and an immunohistochemical account over 14 days. To date, there has been no data published combining both microarray expression and immunohistochemical expression of KGF1, FGF2, or FGF10. By utilizing multiple techniques, insights into where and when growth factors are important in TM wound healing processes can be elucidated. Expression of a gene does not necessarily lead to a translated protein; there may also be a time lag in translation. This research correlates the expression of KGF1, FGF2, and FGF10 with its protein in the tissue.

Materials and methods

The experimental design was identical to a previous experiment investigating the role of Epidermal Growth Factor in the healing TM of rats (Santa Maria et al. 2010b). 410 male Sprague–Dawley rats (Rattus Norvegicus) weighing 250–300 g were obtained from the Animal Resources Centre in Murdoch, Western Australia and housed within the University of Western Australia. The University of Western Australia’s Animal Ethics Committee approved all experimental procedures. A final number of 240 individual rats were used for microarray and 17 for immunohistochemistry. Food and water was provided ad libitum and 12-h light/dark cycles were maintained.

Before perforation, both ears of all rats were examined for signs of inflammation, trauma, or disease. No rats were excluded for these reasons.

Microarray analysis group: rats were randomly selected and placed into either control or perforation groups. Rats were again randomly allocated into time point groups to be sacrificed at 12, 24, 36 h, and at days two through to seven.

Immunohistochemistry group: a rat was allocated to be sacrificed at 12, 24, 36 h and at days two through to 14.

Perforation protocol: Rats in the perforation groups had their left TMs perforated in the upper outer quadrant of the pars tensa using a sterile 23-gauge needle by the first author. TM perforations were created while the rats were under general anaesthesia using an inhalational technique with isoflurane (oxygen and isoflurane mixture of approximately 2 ml/min oxygen and 5 ml/min (induction) or 2.5 ml/min (maintenance) isoflurane). The TM was approached with a 2 mm ear speculum using an operating otomicroscope (Zeiss, Australia).

Surgical protocol: All rats were sacrificed at their allocated time points under general anaesthesia with 2 ml intracardiac pentobarb. The pars tensa of the TM was extracted bilaterally immediately after confirmation of death. To extract the pars tensa, first a postauricular incision was made which was extended to include transection of the ear canal at the bony cartilaginous junction. An incision was then made between the skull base and skin of neck anteriorly allowing exsanguination to occur and provide minimal blood in the field. Soft tissue was dissected off the tympanic bulla using a periosteal elevator and canal skin was removed using a round knife. Using curved needles the pars tensa was dissected from the handle of the malleus and the annulus.

Sample preservation: Samples for microarray analysis were immediately added to sterile tubes containing RNAlater (Ambion Inc, USA), snap frozen, and stored under liquid nitrogen until required. Samples for immunohistochemistry were placed into 4% paraformaldehyde fixative overnight, followed by washing in 1× phosphate buffered saline (PBS), pH 7.4. Tissue processing was performed overnight in a Leica tissue processor, and subsequently embedded in paraffin wax ready for microtomy.

RNA extraction

RNA extraction was performed within 48 h of tissue collection. RNeasy mini kits (Qiagen, Australia) were used according to the manufacturer’s instructions.

Nanodrop/Agilent bioanalyser analysis

All samples were initially screened using a NanoDrop 1000 spectrophotometer (Thermo Scientific, USA) to assess the concentration and chemical purity of Total RNA. Final reassessment of samples before proceeding to the pooling stage was performed using a NanoDrop 1000 (Thermoscientific) and an Agilent 2100 Bioanalyzer. Samples were rejected if they returned an RNA Integrity Number (RIN) of six or less on the Agilent 2100 Bioanalyzer (Schroeder et al. 2006). In cases where a RIN was not obtained because of low RNA concentration, the profile was visually inspected, and if it contained two obvious rRNA peaks similar to other samples in the same time point with a pass RIN, these samples were also passed.


At the point of microarray, there were 18 rats per time point group and 18 controls. Samples were randomly pooled into three groups (biological replicates) per time point. There were 18 individuals per time point i.e. six samples were pooled per group. Sample pooling was performed to ensure that equal amounts of RNA were taken from each sample and three separate pools were made to enable the inclusion of biological variability in the experiment. Pooled groups were then further assessed using a NanoDrop 1000 (Thermoscientific) and also an Agilent 2100 Bioanalyzer. Pooled groups were rejected under the same criteria stated above.

Microarray analysis

The Agilent One-Color Microarray-Based Gene Expression Analysis Protocol (version 1.0.1) was followed, which included use of Agilent’s 4x44 k whole rat genome arrays (40 in total), One Colour RNA Spike-In Kit, cRNA synthesis and amplification, cRNA purification, 17-h hybridization, and scanning using an Agilent DNA Microarray Scanner and Agilent’s Feature Extraction 9.5.3 Software. Forty arrays were scanned which included at least three and maximum of four biological replicates per time point. A total number of 240 samples were included in this study. Overall 393 rats were used as 153 samples failed one of the quality control steps discussed above. Microarray data was performed using GeneSpring GX9 (Agilent, USA). One microarray (within the day six time point) was rejected, as it appeared to be an outlier in the quality control metrics plot produced by GeneSpring. Further analysis was carried out by GeneSpring on genes identified as either present or marginal. Genes with statistically different expression (P < 0.05) between sequential time points were identified using a One Way ANOVA test with the Benjamin and Hochberg multiple testing correction and Tukey Post-hoc test. The gene list was then filtered on criteria of twofold or more change between time points. Further entities of genes were inspected based on areas of interest in wound healing.

Quantitative real time polymerase chain reaction (qRT–PCR)

Quantitative real time PCR using IQ™ SYBR Green Supermix (Bio-Rad, Australia) and an Opticon qPCR thermal cycler (Bio-Rad) was used to confirm the expression levels of β-actin, desmocolin 2, MMP3, Stfa and LCN2. RPLP2 was selected as the most appropriate housekeeping gene according to the geNorm software (Vandesompele et al. 2002). PCR reactions were repeated in triplicate and also repeated in triplicate for each gene. Gene expression levels were calculated using the Pfaffl method (Pfaffl 2001) and normalized to the RPLP2 expression level. Real-time analysis of the relative quantification of gene transcripts confirmed the expression changes seen in the microarray analysis.


Commercially available primary and secondary antibodies were used for immunoflourescent staining of paraffin-embedded sections. Primary antibodies: donkey polyclonal anti-goat KGF1 (Santa Cruz Biotechnology, USA), goat polyclonal anti-rabbit FGF10 (Santa Cruz Biotechnology), goat polyclonal anti-rabbit FGF2 (AbCam, UK). Secondary antibodies: Alexa Fluor-488 conjugated goat anti-mouse, Alexa Fluor-488 conjugated goat anti-rabbit 546, Alexa Fluor-488 conjugated donkey anti-goat (Molecular Probes, USA). Nuclei were counterstained with 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI), (Molecular Probes).


Following tissue processing and paraffin wax embedding, sections were cut at 4 μm and mounted onto positively charged Histobond™ microscope slides (Marienfeld, Germany). Slides were heated overnight at 56°C to allow the sections to adhere and to melt away excess wax. The staining procedure was performed at room temperature (RT) in a light-tight humidified staining box under low light conditions. Briefly, 4 μm sections were hydrated through a series of xylene, absolute ethanol, and graded ethanols to water, then rinsed in 1× phosphate buffered saline (PBS), pH 7.4 containing 0.1% Triton X-100 (PBS-Tx). Sections were incubated in 3% hydrogen peroxide (H2O2) in methanol for 5 min at RT, washed three times in PBS-Tx, then blocked with 20% normal horse serum for 30 min. Incubation with primary antibodies, was performed for one h then washed in three washes of PBS-Tx. Subsequent incubation with the corresponding secondary antibody (1:500), was for 60 min followed by three washes in PBS-Tx. Finally, cell nuclei were counterstained with DAPI for 20 min, washed in three changes of PBS-Tx, cover slipped in an anti-fade mounting medium and sealed with nail varnish.

Confocal laser scanning microscopy and imaging

KGF1 and FGF10

Images were captured sequentially using a BioRad MRC 1000/1024 UV Laser Scanning confocal microscope (Bio-Rad, UK) on Nikon Diaphot 300 (Nikon, Japan) using a dry 20× objective (Nikon Fluor, NA 0.75). A 488 nm blue laser was used to excite AF-488 secondary antibodies and fluorescence emission detected through a 522/35 nm bandpass filter. A UV laser (351 nm) was used for the excitation of DAPI fluorescence emission and detected through a 455/30 nm bandpass filter. Images were saved as 8-bit greyscale images and were subsequently merged with ImageJ (1.42e) (Rasband 1997–2004). All images were saved in TIFF format. Negative controls were run with no primary antibody and these produced no positive staining.


Images were captured sequentially using a Nikon A1Si confocal laser-scanning microscope (Nikon, Japan) on a Nikon TiE (Nikon, Japan) with an oil 40× objective (Nikon Pan Fluor, NA 1.30). The pinhole was set to 1.0 Airy unit (A.U.). A 488 nm blue laser was used to excite AF-488 secondary antibodies and fluorescence emission detected through a 525/50 nm bandpass filter. A 561 nm green laser was used to excite AF-546 secondary antibodies and fluorescence emission detected through a 585/50 nm bandpass emission filter. A violet laser (405 nm) was used for the excitation of DAPI and fluorescence emission detected through a 450/50 nm bandpass filter. Images were saved as 12-bit greyscale images and were subsequently merged using with ImageJ (1.42e), (Wayne Rasband, All images were saved in TIFF format.



The microarray data discussed in this publication is from a larger study, the data of which has been deposited in NCBI’s Gene Expression Omnibus (Edgar et al. 2002) and are accessible through GEO Series accession number GSE17698 (


Two probes where associated with FGF2 and their normalized signal intensities as detected by microarray, in the week following perforation, are shown in Fig. 1. Expression of both probes shows a reduction from 12 to 24 h, and a general increase in expression from day 3 to day 5.
Fig. 1

The normalised signal intensities of KGF1, FGF2, and FGF10 at the time points during 7 days following perforation

Figure 2a presents the protein expression of FGF2 by immunohistochemical staining over the 14 days of wound healing following perforation. The control TM showed no protein expression of FGF2. At 12 h, FGF2 expression was visible on the malleus side of the perforation within the keratinocyte layer. Similar expression within the keratinocyte layer is seen until perforation closure on day six and seven. A wave of new keratinocyte growth is seen migrating toward the perforation site at the 48-h time point, again, emanating from the malleus side. By day seven, with the perforation now closed, FGF2 expression within the keratinocyte layer begins to increase in intensity. Although at day eight and nine, weak expression of FGF2 is visible in the connective tissue layer, the majority of protein expression is continuing to increase in intensity within the keratinocyte layer. No FGF2 extracellular expression was visible within the connective tissue layer before the seven-day time point. However, strong intracellular localization of FGF2 within the connective tissue layer was seen on day five, seven, and eleven. At day 10, FGF2 expression was decreasing within the keratinocyte layer and by day 14 levels resembled that of the control TM.
Fig. 2

a, b Immunofluorescent staining patterns for FGF2 on rat tympanic membrane paraffin sections imaged by laser-scanning confocal microscopy (Bio-Rad). FGF2 positive cells appear green (AF-488) and all cell nuclei, counterstained with DAPI, are visualised in red. The white arrow locates the region of the perforation. Scale bar = 20 μm


The normalized signal intensities of KGF1 detected by transcriptome analysis over the 7 days of wound healing post perforation are displayed in Fig. 1. This shows a peak at 12 h following injury. At 24 h, KGF1 levels return to that seen in the control TM. There was another increase in expression at day three, and continued until day six. Levels began to decline on day seven.

The localization of KGF1 by immunohistochemical staining is shown in Fig. 2b. The peak expression as detected by microarray correlates with the strong KGF1 protein expression visible at 12 h in both the connective tissue and keratinocyte layers. The reduced expression seen at the 24-h time point by transcriptome profiling, correlates with a drop in expression within the connective tissue layer, however KGF1 protein expression persists in the keratinocyte layer until 48 h. KGF1 protein expression remains relatively weak before reappearing in the keratinocyte layer at day seven. This pattern continues and peaks at day nine before decreasing sharply. From day 10 through to 14, expression of KGF1 remains low (Fig. 3a, b).
Fig. 3

a, b Immunofluorescent staining patterns for KGF1 on rat tympanic membrane paraffin sections imaged by laser-scanning confocal microscopy (Bio-Rad). KGF1 positive cells appear green (AF-488) and all cell nuclei, counterstained with DAPI, are visualised in red. The white arrow locates the region of the perforation. Scale bar = 20 μm


The normalized signal intensities of FGF10 detected by transcriptome analysis over the 7 days of wound healing post TM perforation are displayed in Fig. 1. FGF10 expression remained relatively stable over the first 7 days of wound healing, with only a slight reduction in expression following the acute phase of wound healing. This corresponds to the 48-h time point.

Figure 2c shows the immunohistochemical expression of FGF10 over the 14-day period following TM perforation. Despite the unchanging expression of FGF10 shown by microarray analysis, a number of changes in protein expression throughout all time points were visualized by immunohistochemical staining. The TM control tissue exhibited weak protein expression. An increase in FGF10 expression at 36 h was seen on the malleus side of the perforation, localized mainly to the connective tissue layer. This expression continued until day three when it intensified before amassing beneath the keratin layer on day four. FGF10 expression decreased to levels seen in the control tissue by day 5. There is a moderate increase in expression throughout the connective tissue layer at day six. Day seven time point tissue shows a dramatic difference in protein expression between the keratinocyte layer, which is very strong, and the connective tissue layer, which exhibited low to moderate expression. This pattern continues until day 11 when protein expression decreased to appear weak in all layers. Similar expression was present up to and including day 14 time points Fig. 4a, b.
Fig. 4

a, b Immunofluorescent staining patterns for FGF10 on rat tympanic membrane paraffin sections imaged by laser-scanning confocal microscopy (Bio-Rad). FGF10 positive cells appear green (AF-488) and all cell nuclei, counterstained with DAPI, are visualised in red. The white arrow locates the region of the perforation. Scale bar = 20 μm


The benefit of using the rat in research of the tympanic membrane has been well documented. They are similar to the human TM in structure (Schmidt and Hellstrom 1991; Chole and Kodama 1989) and composition (Chole and Kodama 1989; Lim 1968a; Lim 1968b). Rats are readily available worldwide and have the added advantage of easy accessibility. Surgically, transcanal approaches to the rat TM are simple as the whole TM can be viewed from one position via an otomicroscope.


A study in guinea pigs reported that in the 3 days following perforation, FGF2 was expressed in the perforated area, on pericytes and polynuclear cells and has an additional role in stimulating chemotaxis (Mondain and Ryan 1995). FGF2 have been detected in the middle ear mucosa and inflammatory cells surrounding the perforation as early as 24–48 h post injury (Fina et al. 1991). In our study, the transcriptome profile showed a general increase in expression from day three to day five. Other reports indicate that FGF2 mRNA increases gradually following perforation and remain elevated for 5–7 days (Ishimoto and Ishibashi 2002; Ishibashi et al. 1998). The localization of FGF2 was exclusively seen within the keratinocyte layer on the malleus side at 12 h following perforation until closure. This evidence suggests that FGF2 plays a key role in the migration of keratinocytes to close the perforation. The relative inactivity on the annulus side suggests the annulus does not play the same crucial role as the malleus in TM wound healing. Our study mirrored the guinea pig study with FGF2 expression not detectable in normal TM tissue (Mondain and Ryan 1995). The continued FGF2 expression following perforation closure suggests it has a role in more than just the migration of keratinocytes.

FGF2 has been applied to TM perforations in an attempt to improve wound healing. One study reported FGF2 application following perforation might result in greater thickening of the connective tissue layer (Mondain and Ryan 1993) of non perforated areas by up to twenty times (Mondain et al. 1991; Fina et al. 1993). FGF2 and TGFα were found to induce hyperplasia at the proliferation centres of both the epithelial and intermediate layers (not mucosal) but not at the perforation edge (Ishimoto et al. 2002). In one study, macroscopic closure was found to occur slightly earlier and microscopically by day seven there are already non-orientated fibroblasts in the healed area. By day 10 the healed area was reportedly twice as thick as the untreated TMs at the same time point. Orientation of the fibroblast is already underway (Mondain et al. 1991). Another study showed treated TMs healed on average 4 days earlier than control TMs (Vrabec et al. 1994). Further studies showed no significant difference in healing time (Chauvin et al. 1999; Friedman et al. 1997). Compared to untreated TMs, FGF2 treated TM tissue exhibited fibroblast orientation which was more parallel to the TM surface (Mondain and Ryan 1993); (Fina et al. 1991) and greater epidermal proliferation (Lim 1995). FGF2 also induces vasodilation in the TM, possibly explaining the accelerated healing of perforations following application of FGF2 (Mondain and Ryan 1994). Numerous other studies have shown similar results with FGF2 (Fina et al. 1993; Chauvin et al. 1999; Friedman et al. 1997; Kato and Jackler 1996). The functional outcome between FGF2 treated perforations and control ears, as measured by tympanometry, is the same (Vrabec et al. 1994). When neutralizing antibody to FGF2 is applied to TM perforations, the result is fewer epithelial and mononuclear cells (Mondain et al. 1995). A clinical study reported in Chinese, cited by Ma et al. (2002), using FGF2 on TM perforations showed a significant benefit over placebos in perforation closure.


Most cells in a wounded area synthesize FGF, however only KGF1 is shown to be synthesized by fibroblasts. KGF1 stimulates mitosis in keratinocytes and not fibroblasts (Finch et al. 1989). It stimulates the production of proteinases (e.g. Plasminogen activator) that are involved in tissue remodelling and growth factor activation (Clymer et al. 1996). Studies in the TM following perforation have reported KGF1 mRNA expression to be induced early, peaking 3 days after wounding and then declining (Ishibashi et al. 1998; Ishimoto and Ishibashi 2002). This study suggests the KGF1 has a role in the acute inflammatory process with an up regulation at 12 h. There is also confirmation that a second peak at day three of KGF1 expression promoted differentiation and organization of healing rather than rate of repair (Clymer et al. 1996). The main role of KGF1 in TM wound healing may only be in the closure of the perforation as the following time points show both a decline in expression and a reduction in staining.

Studies applying KGF1 to TM perforations suggest it promotes differentiation and organisation of healing rather than rate of repair (Clymer et al. 1996). Topical application of KGF1 does not enhance the closure rate of TM perforation but the repair process is reportedly more organised, showing greater hyperplasia and a thickened healed area (Clymer et al. 1996). When KGF1, FGF2, and TGFa were applied to perforations in the setting of systemic glucocorticoid treatment, only KGF1 improved keratinocyte cell migration.


There have been no previous publications implicating the role of FGF10 in the healing of the TM. FGF-10 (also known as KGF 2) is highly homologous to KGF1 (Marchese et al. 2001) and has similar receptor binding properties and target cell specificities, but is differentially regulated by components of the extracellular matrix (Braun et al. 2004). Despite this, this study did not show a correlation of FGF10 to KGF1 suggesting different roles in TM wound healing. There is no expression of FGF10 in control TM tissue. Once the TM is perforated, like KGF1, FGF10 can be seen on the malleus side of the perforation. Unlike KGF1, FGF10 expression was predominantly localized to the connective tissue layer suggesting its site of action is here rather than the keratinocytes. Only on day seven with the perforation closed, was FGF10 predominantly found within the keratinocyte layer. Additionally, unlike KGF1, FGF10 expression persisted in time points post day seven following closure. This suggests an additional role for FGF10 beyond perforation closure.

Topically applied FGF10 in an incisional wound healing model reported that treatment resulted in improvement of wound healing as characterized by an increase in breaking strength, collagen content, and epidermal thickness (Jimenez and Rampy 1999). FGF10 is similar to KGF1 as a potent mitogen for human keratinocytes, but it also promotes the expression of both early (keratin 1, keratin 10) and late (filaggrin) differentiation markers in response to Ca2+ signalling, and seems to sustain proliferative activity in suprabasal stratified cells. Like KGF1, FGF10 is able to induce tyrosine phosphorylation of keratinocyte growth factor receptor and of cellular substrates such as PLCgamma (Marchese et al. 2001).


KGF1, FGF2, and FGF10 all play important key roles in TM wound healing. The role of KGF1 and FGF2 is primarily related to the proliferation and migration of keratinocytes into a wound site. KGF1’s role appears to be limited to this task, while FGF2 has an ongoing link with keratinocytes in the 7 days following perforation. FGF10, whilst being homologous to KGF1, has a different role in TM wound healing. FGF10’s role is predominantly in the connective tissue layer.


Garnett Passe and Rodney Williams Memorial Foundation for the funding of this study. Assoc Professor Robert Eikelboom for editorial support. Medtronic for sponsorship and supply of microsurgical instruments. Zeiss for sponsorship of the otomicroscope. The authors acknowledge the facilities, scientific and technical assistance of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterisation & Analysis, The University of Western Australia, a facility funded by The University, State and Commonwealth Governments.

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