Mapping proteolytic cancer cell-extracellular matrix interfaces
- First Online:
- Cite this article as:
- Wolf, K. & Friedl, P. Clin Exp Metastasis (2009) 26: 289. doi:10.1007/s10585-008-9190-2
For cancer progression and metastatic dissemination, cancer cells migrate and penetrate through extracellular tissues. Cancer invasion is frequently facilitated by proteolytic processing of components of the extracellular matrix (ECM). The cellular regions mediating proteolysis are diverse and depend upon the physical structure, composition, and dimensionality of the ECM contacted by the cell surface. Cancer cells migrating across 2D substrate contain proteolytic structures such as lamellipodia, invadopodia, and the trailing edge. Likewise, invasive mesenchymal migration through 3D fibrillar ECM, as monitored for HT1080 fibrosarcoma and MDA-MB-231 breast carcinoma cells by submicron-resolved imaging, is mediated by several types of proteolytic structures rich in filamentous actin, ß1 integrin, and MT1-MMP with distinct location and function. These comprise (i) anterior pseudopod bifurcataions and the nucleus corresponding to zones of local cell compression by constraining collagen fibers, (ii) lateral small spikes that protrude into the ECM and cause small spot-like proteolytic foci, and (iii) a strongly proteolytic trailing edge sliding along reorganized ECM fibers. Through their combined action these proteolytic surface structures cleave, remove, and realign ECM barriers, support rear end retraction, generate tube-like matrix defects and laterally widen existing tracks during 3D tissue invasion.
KeywordsCancerInvasionMigrationProteasesProteolysisECMLamellipodInvadopodPseudopodLateral spikeCell compressionMicrotrack
Focal adhesion kinase
Green Fluorescent Protein
Membrane-type 1 MMP
Urokinase plasminogen activator
Urokinase plasminogen activator receptor
The progression of cancer is a complex process that includes the migration-associated proteolytic interaction of cancer cells with extracellular matrix (ECM). Cell migration requires anterior protrusion, attachment, localized ECM degradation together with acto-myosin contraction and rear end detachment . In vivo, invading cells are confronted with structurally distinct types and spatial dimensions of tissue structures, such as two-dimensional (2D) surfaces or three-dimensional (3D) ECM.
3D tissue structures in vivo display a heterogeneous network of fiber scaffolds of variable density, orientation, and mechanic strength. Connective tissues rich in fibrillar collagen I or III, elastic fibers and other ECM components are present in and around most organs, such as skin, breast or gut. Cancer cells need to penetrate such interstitial tissue before reaching vessels for hematogenic or lymphogenic dissemination [5–7]. While invading, tumor cells are surrounded by ECM that simultaneously acts as structural barrier. Therefore, proteolytic cell structures arise from all sides of the cell, interact with 3D ECM circumferentially, and support local disruption and restructuring of the tissue scaffold (Fig. 1c). In 3D invasion, proteolytic cell surface structures are homologous in subcellular position and molecular regulation to those formed in 2D models, albeit their precise topography and mode of ECM remodelling are different. As an example, actin-rich flat lamellipodia formed on 2D substrate correspond to cylindrical-shaped pseudopodia that extend along and protrude through fibrillar tissue in a 3D environment (Fig. 1) [8, 9].
Here we visualize and classify the subcellular location and function of proteolytic structures in migrating cancer cells. Known cellular regions degrading 2D surfaces will be compared to newly identified surface-localized structures executing 3D ECM breakdown.
Experimental models for imaging protease function at the cell-ECM interface
Experimental models for examining protease activity and ECM degradation in vitro and in vivo
Loss of substrate
Thin layer of unpolymerized or polymerized ECM coated onto 2D carrier (Substrate: gelatin, collagen, fibronectin, matrigel, dentin, apatite; carrier: glas, plastic); detection of degradation by using fluorescently labelled ECM components [28, 37, 60]
Basal lamina-like membrane produced by cells on collagen 
In vitro reconstituted 3D ECM polymers (gelatin, collagen, matrigel, fibrin, functionalized hydrogels)
Layer of unpolymerized or polymerized substrate (collagen, gelatin, casein, matrigel) containing quenched fluorophore
DQFITC-collagen I ,
Polymerized 3D ECM (collagen I and IV), containing quenched fluorophore
DQFITC-collagen I ,
DQFITC-collagen IV 
Detection of cleavage-site specific epitopes
Isolated decellularized basement membrane from in vivo tissue 
Proteolytic structures during migration on 2D surfaces
Molecular contents of proteolytic cell structures and associated cell track of migrating cells
Cell front/mid body
Proteolytic structures during 3D tissue invasion
Until recently the cell structures generating surface-associated proteolysis in 3D tissues were unknown. However, novel multimodal imaging technology, such as confocal fluorescence and reflection, multiphoton-excited fluorescence and second harmonic detection, or scanning electron microscopy, now permits the reconstruction of proteolytic cell-matrix interactions in time and space. When embedded within 3D ECM, such as fibrillar fibrin or collagen lattices, or when seeded onto a 3D dermis slice ex vivo, many cell types including fibroblasts and cancer cells polarize and start to migrate [7, 9, 22, 30–32]. Cell surface adhesion receptors such as integrins, and ECM-degrading proteases can focally interact with the substrate in different cellular locations that convey specific functions, including (i) the leading edge and protruding pseudopodia with anterior zones of ECM degradation, (ii) the mid region that glides along the substrate and cleaves lateral ECM structures, and (iii) the retracting trailing edge that weakens adhesion bonds and supports forward movement of the cell (Fig. 1c) . As on 2D surfaces, proteolytic cellular zones in migration through 3D ECM contain filamentous actin, actin-associated adaptor proteins, and adhesion receptors in close proximity with surface proteases (components listed in Table 2).
Proteolytic structures at the leading edge
Compression zones along the mid cell body
To explore the spatio-temporal fate of such processed fibers by live cells, a collagen lattice labelled with quenched FITC molecules was used for dynamic imaging of collagenolysis [9, 41]. As the trailing edge moves forward (Fig. 3c, asterisk), a compression belt (empty arrowhead) becomes proteolytically opened which allows expansion of the cell diameter by 2 μm (Fig. 3c, white arrowhead). Thus, at regions of cellular compression, collagen fibers become sequentially degraded and opened to generate space for the migrating cell body and nucleus and are realigned in the direction of forward movement (Fig. 2c, white arrows) .
Trailing edge and microtrack formation
Whereas these steps of ECM remodelling form a continuous cycle that is embedded within the cell migration cycle, each protease-containing actin-rich structure is morphologically distinct and mediates a particular function. First, the tips of growing pseudopodia attach to the substrate, mediate integrin-dependent traction force, and bring the protease in close contact to the substrate thus targeting MMPs towards ECM substrate. Belt-like cell compressions by sterically impeding ECM structures cause locally confined cleavage, preferentially of those fibers that arrest pseudopod bifurcations or the nucleus. Thus, local pressure at regions of physical resistance against the cell body elicit surface protease activity by as yet unknown mechanisms. Next, lateral spikes and pseudopodia which do not contribute to pulling force generation push outward towards encountered scaffold structures, probe and focally degrade the ECM at their tips, and rapidly retract. Their function may include additional anchoring and stabilization of the cell body within the surrounding ECM and additional ECM processing. Although the proteolytic amount per spike is only minor, a sufficient spike frequency may substantially contribute to lateral ECM modification. Third, the trailing edge, in contrast to other proteolytic zones, is neither protrusive nor confronted with local pressure. Rather a relatively large, diffuse zone of protease distribution and ECM degradation is being formed, suggestive of constitutive and substantial ECM remodelling and ECM fragment generation. Track formation is therefore a consequence of the combined action of distinct proteolytic zones formed during cell migration allowing the cell to cleave and shape ECM fibers into a biochemically altered, parallel oriented cell track. The emerging path might be additionally modulated by ECM degradation fragments from laminin, fibronection or collagen that were shown to act chemotactically on neighbouring cells [51–53]. Microtrack-deposited proteases may cleave and thereby activate ECM-bound growth factors and chemokines produced by tumors or tumor-associated cells such as fibroblasts and macrophages and establish a chemotactic gradient [54, 55]. All these structural and biochemical ECM alterations and new contents could attract tumor cells into an already established track and thereby mediate their conversion to multicellular invasion [9, 50].
Whereas these studies show the ECM remodelling during invasive cell migration in 3D model matrix, further work is needed to show proteolytic cell migration in vivo, i.e. the location and function of each proteolytic cell structure in the context of even more structural complex tissues. To visualize such proteolytic processes in vivo [22, 56], the development of protease-specific quenched substrates is required that not only freely reach the site of interest but, after cleavage, adopt a non-diffusive state with high temporal and spatial fidelity by, e.g. precipitation in situ. Together, high-resolution microscopy of cell invasion in vitro and in vivo will provide a detailed map of different proteolytic cell structures and their specific function in a tissue and organ context.
We thank M. Ott for excellent technical assistance; S. Stack and Y. Wu as well as E. Deryugina and A.Y. Strongin for providing MDA-MB-231 and HT-1080 cell lines, respectively; D. Pei for supply of human MT1-MMP-GFP cDNA; and L. King (IBEX, Canada) for supply of COL2¾Cshort antibody. This work was supported by the Deutsche Forschungsgemeinschaft (FR 1155/7-1) and the Deutsche Krebshilfe (AZ 106950).
This article is distributed under the terms of the Creative Commons Attribution Noncommercial License which permits any noncommercial use, distribution, and reproduction in any medium, provided the original author(s) and source are credited.