A new procedure for rapid, high yield purification of Type I collagen for tissue engineering
Introduction
Collagen is a major scaffold for biotechnological applications, e.g. in tissue engineering. This is one of the reasons why the molecular mechanism for the biosynthetic assembly of collagen is of high interest. Collagen is the major component in the extracellular matrix and Types I, II and III are the most abundant collagens which form fibrils responsible for tensile strength [1], [2]. Due to its high accessibility and compatibility, Type I collagen is one of the most well-studied collagens and is already widely used as a bioscaffold in medicine and cell biology [3], [4], [5], [6]. Type I collagen is trimeric [(α1)2α2] and exists as triple helix. There are several common sources for collagen: natural tissues such as skin and tendons [7], [8], synthetic collagen peptides [9], [10] and cell cultures [11]. The spontaneous formation of triple helix from isolated α1 and α2 polypeptides in vitro has been studied extensively [12], [13], [14], [15] and recent studies using synthetic model peptides [9], [10] show the self-association or self-assembly of these peptides to a native-like triple-helix structure characteristically found in collagen fibrils.
To isolate the collagen from cell culture or natural tissues the two most commonly used methods are neutral salt-extraction and low concentration acid-extraction using acetic acid and citric acid at a concentration of 0.05–0.5 M [16], [17]. Reports that Type I collagen and procollagen have also been extracted using 8–10 M urea also exist [18], [19]. However, this procedure has been described to yield only poor quality material, with significant degradation [18], making it non-attractive for further development.
Collagen has been successfully applied for different purposes, e.g. as drug delivery system [20] and scaffold in tissue engineering [21], [22]. In comparison to other biomaterials, naturally derived collagen shows excellent biocompatibility and safety due to its high abundance in all vertebrate animals and high biodegradability [23]. In addition, collagen also exhibits very low antigenicity [24].
Fibroblasts cultured using collagen matrix containing an ordered 3D-structure (3D cell derived matrix) take on a spindle-shaped morphology, similar to in vivo fibroblast morphology. These cells show also increased proliferation, migration and adhesion compared to fibroblasts cultured on a collagen matrix without ordered structure (2D flattened cell-derived matrix) [25]. Binding of collagen to the integrins and other cellular receptors mediates cell adhesion and regulates the cell motility. Moreover, the interaction with proteoglycans and other components in extracellular matrix (ECM) can regulate the mechanical properties of different tissues [15], [26]. Cell behavior can be affected by the changes of the ECM, in particular the fibrous collagen. It was shown that changes in the compression and alignments of the fibrils of collagen can lead to cellular signaling resulting in the formation of a regular geometric system pattern characteristic for the respective tissue [27]. Therefore, the matrix used in tissue engineering may be crucial for constructing a defined tissue- or organ-like structure.
In this study we show that extraction of rat tail tendon with 9 M urea yields large quantities of homogeneous Type I collagen (α1, α2). The homogeneity of the preparation was confirmed by mass spectrometry (MS) and amino acid composition analysis. The morphology of fibroblasts grown on urea-extracted collagen (UC) and acetic acid/extracted collagen (AC) differed clearly. This was reflected in studies on a transcriptional level using qRT-PCR, whereby the regulation of Fak1, Rac1, MMP9 COL1A1 was determined. Our results indicate that purified urea-extracted collagen results in higher motility and reduced stress levels of fibroblasts than acetic acid-extracted collagen showing the suitability of urea-extracted collagen for biotechnological applications and tissue engineering.
Section snippets
Protein isolation from rat tail tendons (RTTs)
Rat tail tendon (RTT), teased out from 3-month-old rats was washed extensively in phosphate saline buffer (PBS) pH 7.4 at 4 °C. In order to get rid of traces of isocyanic acid in urea, the urea used was purified prior to extraction by an incubation with Biorad AG501-X8 resine (Nr.: 142424) for 30 min. The tendon was extracted with 9 M urea at 25 °C for 20 h, followed by a centrifugation at 4000 × g for 30 min to sediment the insoluble components. The urea-soluble proteins were purified by
Collagen extraction and SDS-PAGE analysis
Collagen samples obtained from rat tail tendon of 3-month-old rats after extraction with 9 M urea following our procedure yielded a clear solution. The yield of the urea-extraction without purification via gel-filtration was calculated from three isolations using different amounts of rat tail tendons. To determine the yield, the tendons were first washed in PBS and water then lyophilized. The dry weight of the lyophilized rat tail tendon was determined and after 20 h incubation in urea (see
Discussion
We have established a fast two-step urea-extraction procedure to isolate and purify Type I collagen from rat tail tendon. Although the extraction of collagen Type I using urea was reported more than 30 years ago [18], [42], [43], this method has been limited in its application maybe due to reports that only low quality collagen can be obtained with this method. In our study, we found that the collagen isolated using 9 M ultrapure urea is of high quality and shows a reversible aggregation
Acknowledgements
We would like to thank Mrs. M. Riedl and Dr. H. Weber for the technical support. We also thank Dr. W. Haehnel for kindly providing mass spectrometry facilities. We thank Ms. C. Autenrieth for performing the Gaussian analysis. Especially, we want to thank Prof. A. Veis for fruitful discussions. This work was supported by a fellowship of the Peter und Traudl Engelhorn Foundation to X.X.
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Present address: Institute of Experimental Internal Medicine, Medical Faculty, Otto von Guericke University, Leipziger Str. 44 39120 Magdeburg, Germany.