Matrix Metalloproteinase Biology E-Book
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- Herausgeber: John Wiley & Sons
- Kategorie: Wissenschaft und neue Technologien
- Sprache: Englisch
Discussing recent advances in the field of matrix metalloproteinase (MMP) research from a multidisciplinary perspective, Matrix Metalloproteinase Biologyis a collection of chapters written by leaders in the field of MMPs. The book focuses on the challenges of understanding the mechanisms substrate degradation by MMPs, as well as how these enzymes are able to degrade large, highly ordered substrates such as collagen. All topics addressed are considered in relation to disease progression including roles in cancer metastasis, rheumatoid arthritis and other inflammatory diseases.
The text first provides an overview of MMPs, focusing on the history, the development and failures of small molecule inhibitors in clinical trials, and work with TIMPS, the endogenous inhibitors of MMPs. These introductory chapters establish the foundation for later discussion of the recent progress on the design of different types of inhibitors, including novel antibody based therapeutics. The following section emphasizes research using novel methods to further the study of the MMPs. The third and final section focuses on in vivo research, particularly with respect to cancer models, degradation of the extracellular matrix, and MMP involvement in other disease states.
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Veröffentlichungsjahr: 2015
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Table of Contents
Cover
Title Page
Copyright
List of Contributors
Chapter 1: Matrix Metalloproteinases: From Structure to Function
1.1 Introduction
1.2 Structures of MMPs
1.3 Overview of MMP substrate specificity
1.4 Selective mechanisms of action
Acknowledgments
References
Chapter 2: Dynamics and Mechanism of Substrate Recognition by Matrix Metalloproteases
2.1 Introduction
2.2 Conformational flexibility of MMPs is inexorably linked to collagen proteolysis
2.3 Dynamics of MMP-2 and MMP-9 interaction with gelatin
2.4 Surface diffusion: a common mechanism for substrate interaction adapted by MMP-2 and MMP-9
2.5 Dynamics of MMP interaction with collagen fibrils
2.6 Mechanism of interaction of MMP-1, MMP-2, MMP-9, and MMP-14 with collagen substrate involves surface diffusion
2.7 Mechanism of MMP-1 diffusion on native collagen fibrils
2.8 Triple helical collagen cleavage–diffusion coupling
2.9 Conclusions
References
Chapter 3: Matrix Metalloproteinases: From Structure to Function
3.1 Introduction
3.2 Classification and structural features
3.3 Catalytic mechanism
3.4 Intra- and inter-domain flexibility
3.5 Elastin and collagen degradation
References
Chapter 4: Metzincin Modulators
4.1 Inhibitors
Summary and future directions
References
Chapter 5: Therapeutics Targeting Matrix Metalloproteinases
5.1 Introduction
5.2 Peptidomimetic MMP inhibitors
5.3 Structure-based MMPI drug design
5.4 Mechanism-based MMPI design
5.5 Allosteric MMPI design
5.6 Macromolecular MMP inhibitors
5.7 Chemically-Modified tetracyclines
5.8 Alternative approaches
5.9 MMPs as anti-targets
5.10 Conclusions
References
Chapter 6: Matrix Metalloproteinase Modification of Extracellular Matrix-Mediated Signaling
6.1 Introduction
6.2 The extracellular matrix as a source for signaling ligands
6.3 ECM and mechanosensory signal transduction
6.4 Matrix remodeling and modification of mechano-sensory signaling
6.5 Conclusions and future directions
References
Chapter 7: Meprin and ADAM Metalloproteases: Two Sides of the Same Coin?
7.1 Introduction
7.2 Meprin metalloproteases
7.3 Structure of meprin α and meprin β
7.4 Proteomics for the identification of meprin substrates
7.5 Meprins in health and disease
7.6 Proteolytic back-and-forth of meprins and ADAMs
7.7 Collagen fibril formation
7.8 Angiogenesis and cancer
7.9 Inflammation
7.10 ADAM Proteases
7.11 The ADAM family of proteases
7.12 Orchestration of different pathways by ADAM17
7.13 Regulation of ADAM17 activity
7.14 Role of ADAM17
in vivo
7.15 Role of ADAM17 in humans
References
Chapter 8: Subtracting Matrix out of the Equation: New Key Roles of Matrix Metalloproteinases in Innate Immunity and Disease
8.1 The tale of a Frog's tail
8.2 The MMP family
8.3 Making the cut as immune regulators
8.4 Enter the “omics” era: genomics, proteomics and degradomics
8.5 ECM versus Non-ECM MMP substrates
8.6 Moonlighting protein substrates: intracellular proteins cleaved outside the cell
8.7 Intracellular protein substrates cleaved inside the cell by MMPs
8.8 Non-proteolytic roles of MMPs: missed in the myth?
8.9 The fairy TAIL of a frog Has an unexpected ending
Acknowledgements
References
Chapter 9: MMPs: From Genomics to Degradomics
9.1 Introduction
9.2 Degradomics – an overview
9.3 Conclusions
Acknowledgments
References
Chapter 10: MMPs in Biology and Medicine
10.1 Introduction
10.2 Functional roles of MMPs and ADAMs
10.3 MMPs as diagnostic and prognostic biomarkers of cancer
10.4 MMPs/ADAMs as diagnostic and prognostic biomarkers for non-neoplastic diseases
10.5 MMPs as biomarkers of therapeutic efficacy
10.6 MMP-specific molecular imaging for noninvasive disease detection
10.7 Conclusions
Acknowledgments
References
Index
End User License Agreement
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Guide
Cover
Table of Contents
Begin Reading
List of Illustrations
Chapter 1: Matrix Metalloproteinases: From Structure to Function
Figure 1.1 General domain organization of MMPs.
Figure 1.2 Typical structure of the CAT domain of MMPs. Characteristic structural elements are highlighted with arrows. Figure generated using MMP-8 structure (PDB 2OY2) [4].
Figure 1.3 Mechanism of proteolysis catalyzed by MMPs. (Figure prepared based on mechanism proposed by Lovejoy et al. [5]).
Figure 1.4 Fibronectin type II-like module structure and organization. (a) General orientation of FN2 modules of MMP-2. (b) Top view of FN2 modules. Figure prepared using MMP-2 structure (PDB 1CK7) [8].
Figure 1.5 Comparison of MMP linker lengths and sequences. Table was generated after alignment of human MMPs using sequences from the Uniprot database [19] and SeaView 4 [20] and Jalview [21] programs.
Figure 1.6 Typical structure of the HPX domain. The propeller-like structure is composed of four blades (I-IV) and stabilized by a single disulfide bridge, designated with an arrow. In the central tunnel, up to four different ions have been identified (here Ca
2+
is orange and Cl
−
is yellow). This Figure was generated using the HPX domain of MT1-MMP (PDB 3C7X) [23].
Figure 1.7 Structure of TM domain and cytoplasmic tail (residues 518–582) of human MT1-MMP generated by homology modeling [50, 51].
Figure 1.8 Mechanism of the initial steps of collagenolysis. (a) Closed (left) and open/extended (right) forms of MMP-1 in equilibrium. (b) The extended protein binds THP chains 1T-2T at Val23-Leu26 with the HPX domain and the residues around the cleavage site with the CAT domain. The THP is still in a compact conformation. (c) Closed FL-MMP-1 interacting with the released 1T chain (in magenta). (d) After hydrolysis, both peptide fragments (C- and N-terminal) are initially bound to the active site. (e) The C-terminal region of the N-terminal peptide fragment is released. (Reprinted with permission from [16]. Copyright (2012) American Chemical Society).
Chapter 3: Matrix Metalloproteinases: From Structure to Function
Figure 3.1 Structural organization of human MMPs with the corresponding linker length.
Figure 3.2 Ribbon representation of the inactive human proMMP-2. The prodomain, catalytic domain, fibronectin domains, and hemopexin domain are shown in yellow, red, blue, and orange, respectively. The catalytic and the structural zinc ions are represented as magenta spheres and calcium ions as green spheres.
Figure 3.3 Stereo view of the catalytic (a) and hemopexin-like (b) domains of MMP-12 represented as ribbons. In the catalytic domain α-helices, β-strands, and loops are organized in a L1-β1-L2-α1-L3-β2-L4-β3-L5-β4-L6-β5-L7-α2-L8-α3 topology. The catalytic (Zn1) and the structural (Zn2) zinc ions are shown as magenta spheres of arbitrary radius. The first (Ca1), the second (Ca2), the third (Ca3) calcium ions and the calcium ion in the hemopexin-like domain are shown as blue spheres. The three histidines that bind the catalytic zinc and the catalytically relevant glutamate are represented as cyan sticks. Strands and helices are labeled with numbers and greek letters. The hemopexin-like domain is constituted by four β-sheets of four antiparallel β-strands that folds in a symmetric four-blade propeller [53, 67]. The central deep tunnel filled by water molecules is closed by a calcium ion (Ca4) at the bottom.
Figure 3.4 Proteolysis of the collagen fragment ProGlnGlyIleAlaGly by MMP-12. (a) Active site of the free enzyme before the interaction with the substrate. (b) Calculated model of the gemdiol intermediate. (c) X-ray structure of the two-peptide intermediate obtained by soaking the active uninhibited MMP-12 crystals with the collagen peptide. (d) Adduct of MMP-12 with the peptide fragment IleAlaGly after the release of the C-terminal fragment.
Figure 3.5 Pattern of residues interacting with elastin fragments in the isolated catalytic and hemopexin-like domains (a) and in the full length protein (b). The larger effects observed in the full length protein suggest cooperativity of the two domains in binding of elastin fragments.
Figure 3.6 Closed (left) and open/extended (right) forms of FL-MMP-1 in equilibrium. The catalytic zinc ion is represented as a magenta sphere.
Figure 3.7 Proposed mechanism for collagenolysis. In panel (a), from the top (the experimentally-driven docked complex between FL-MMP-1 and THP) to the bottom (the unwounded THP bound to the X-ray closed conformation of FL-MMP-1) the intermediate and energetically possible structures generated by HADDOCK [112] to provide a smooth conformational transition between the initial and final states. In panel (b), starting from the experimentally-driven docked complex between FL-MMP-1 and THP (top), the closed FL-MMP-1 interacting with the released 1T chain (in red), the hydrolysis of the 1T chain with both peptide fragments still in place, and the complex with the C-terminal region of the N-terminal peptide released from the active site (bottom).
Figure 3.8 Interaction of FL-MMP-1 with the substrate. In the panel, from the top to the bottom: (a) structure with the highest MO, (b–c) two morphing intermediate steps, (d)the experimentally-driven docked complex where the hemopexin-like domain and the catalytic domain bind the triple-helical collagen. The structure with the highest MO e morphing structures were aligned to the hemopexin-like domain of the docked complex. FL-MMP-1 and THP are represented as white and yellow surfaces, respectively. In blue is the MMP consensus sequence HE
XX
H
XX
G
XX
H and the cleavage site (
Gly-Ile
) in the first chain of THP. The catalytic zinc ion is represented as an orange sphere. To facilitate visualizing the movement of the catalytic domain with respect to the hemopexin-like domain, the blue and red arrows indicate the direction of helices hA and hC of the catalytic domain defined by residues 130–141 and 250–258, respectively.
Chapter 4: Metzincin Modulators
Figure 4.1 Schematic representation of the ADAM17 proteins used in the present study. (a) Full-length ADAM17 (amino acids 1–824). (b) Soluble forms of the cysteine-rich domain (DE, amino acids 476–642), the disintegrin-like domain (D, amino acids 476–580) and the membrane proximal cysteine-rich extension (E, amino acids 581–642) expressed in
E
.
coli
. (c) ADAM17_DE consisting of the cysteine-rich domain (DE) followed by the transmembrane region (TM) of human ADAM17 (amino acids 475–694). Pro, pro-domain; CD, catalytic domain; TM, transmembrane region; IR, intracellular region. (Reproduced with permission from Yamamoto K., Trad A., Baumgart A., Huske L., Lorenzen I., Chalaris A., Grotzinger J., Dechow T., Scheller J., and Rose-John S. (2012) A novel bispecific single-chain antibody for ADAM17 and CD3 induces T-cell-mediated lysis of prostate cancer cells.
Biochem J
, 445 (1) 135-144. © the Biochemical Society).
Figure 4.2 Design and expression of ADAM17-specific A300E-BiTE. Schematic representation of generation of A300E-BiTE. To identify cDNA sequences of V
H
and V
L
of mouse monoclonal antibody, ten primer sets and seven primer sets were used to amplify V
H
and V
L
cDNA. After analysis of DNA sequences from V
H
and V
L
fragments the construct of A300E-scFv and BiTE were introduced into pET23a and pcDNA3.1 vectors respectively. Linker indicates flexible linker (Gly4Ser). The c-Myc and His6 tags are fused for detection and purification respectively. (Reproduced with permission from Yamamoto K., Trad A., Baumgart A., Huske L., Lorenzen I., Chalaris A., Grotzinger J., Dechow T., Scheller J., and Rose-John S. (2012) A novel bispecific single-chain antibody for ADAM17 and CD3 induces T-cell-mediated lysis of prostate cancer cells.
Biochem J
, 445 (1) 135–144. © the Biochemical Society).
Figure 4.3 Experimental overview. (a) The human TACE ectodomain consists of an amino-terminal metalloprotease catalytic domain (light red) and a carboxyl-terminal noncatalytic Dis-Cys domain (light blue) (I-TASSER model). We exploited this multidomain topology to develop a truly specific ADAM inhibitor using two-step antibody phage display. (b) (i) First, the catalytic site of TACE ectodomain was blocked during primary antibody phage-display selections using the small-molecule inhibitor CT1746. This prevented the selection of antibodies with catalytic-cleft epitopes that could cross-react with non-target metalloproteases. (ii) Primary screening revealed the inhibitory scFv antibody clone D1. This scFv bound specifically to the TACE Dis-Cys domain through its variable heavy (V
H
) domain. (iii) A D1-V
H
-bias antibody phage display library was produced to introduce new variable light (neo-V
L
) chains while maintaining the TACE specificity provided by the D1-V
H
. Secondary selections were performed in the absence of CT1746 in order to provide the neo-V
L
chains with uninterrupted access to the TACE catalytic site. (iv) Secondary screening identified several neo-VL scFvs capable of binding the isolated TACE catalytic domain. Due to Dis-Cys domain binding through the D1-V
H
these “cross-domain” antibodies maintained their strict specificity for TACE. D1-V
H
-neo-V
L
scFv clone A12 (D1(A12)) exhibited the highest affinity for the TACE ectodomain and is the most selectively potent cell-surface ADAM inhibitor ever described. (Reproduced with permission from Tape, C. J., Willems, S. H., Dombernowsky, S. L., Stanley, P. L., Fogarasi, M., Ouwehand, W., McCafferty, J., and Murphy, G. (2011) Cross-domain inhibition of TACE ectodomain
Proc Natl Acad Sci
U S A 108, 5578–5583).
Figure 4.4 Collagen-based, peptidomimetic hydroxamates. (Reproduced with permission from Fisher, J. F., and Mobashery, S. (2006) Recent advances in MMP inhibitor design.
Cancer Metastasis Rev
25, 115–136. Copyright © 2006, Springer).
Figure 4.5 Nomenclature used for enzyme and substrate subsites. The arrow marks the site of protease hydrolysis. (Reproduced with permission from Lauer-Fields, J., Brew, K., Whitehead, J. K., Li, S., Hammer, R. P., and Fields, G. B. (2007) Triple-helical transition state analogues: a new class of selective matrix metalloproteinase inhibitors.
J Am Chem Soc
129, 10408–10417).
Figure 4.6 Sequence of triple-helical peptide containing phosphinate group.
Figure 4.7 A comparison of disulfide topology and sequences of human N-TIMP-1 and sarafotoxin 6b. (Adapted from Lauer-Fields, J. L., Cudic, M., Wei, S., Mari, F., Fields, G. B., and Brew, K. (2007) Engineered sarafotoxins as tissue inhibitor of metalloproteinases-like matrix metalloproteinase inhibitors.
J Biol Chem
282, 26948–26955. Rights holder: AMERICAN SOC FOR BIOCHEMISTRY & MOLECULAR BIOLOGY).
Figure 4.8 Inhibition of MMP-13 by 30 different compounds, as monitored by RP-HPLC and fluorescence spectroscopy. The change in RP-HPLC peak areas or relative fluorescence units for 10 nM MMP-13 hydrolysis of 10 μM fTHP-15 or 5 μM Knight fSSP was monitored at an inhibitor concentration of 100 μM. Assays were performed in triplicate. (Reproduced with permission from Lauer-Fields, J. L., Minond, D., Chase, P. S., Baillargeon, P. E., Saldanha, S. A., Stawikowska, R., Hodder, P., and Fields, G. B. (2009) High throughput screening of potentially selective MMP-13 exosite inhibitors utilizing a triple-helical FRET substrate.
Bioorg Med Chem
17, 990–1005. © PERGAMON).
Figure 4.9 Lineweaver–Burk plot of MMP-13 inhibition of fTHP-15 hydrolysis by compound 20 (a) or 24 (b). (Reproduced with permission from Roth, J., Minond, D., Darout, E., Liu, Q., Lauer, J., Hodder, P., Fields, G. B., and Roush, W. R. (2011) Identification of novel, exosite-binding matrix metalloproteinase-13 inhibitor scaffolds.
Bioorg Med Chem Lett
21, 7180–7184. © PERGAMON).
Figure 4.10 β-Gal-(1→3)-GalNAc (TF antigen) and
N
-acetylglucosamine (GlcNAc) found on TNFα and IL6-R.
Figure 4.11 Results of the pilot “scaffold ranking” screen of TPIMS drug-like library against ADAM10 and 17. Shown is an ADAM10 (a) and ADAM17 (b) screen using glycosylated (red checked bars) and non-glycosylated substrate (blue bars). The arrow indicates library containing potential exosite inhibitors of ADAM17. All assays were performed in triplicate. Activity and selectivity of all libraries were confirmed in reversed-phase HPLC-based assays. (c), basic scaffold of library 1344. (Adapted from Minond, D., Cudic, M., Bionda, N., Giulianotti, M., Maida, L., Houghten, R. A., and Fields, G. B. (2012) Discovery of novel inhibitors of a disintegrin and metalloprotease 17 (ADAM17) using glycosylated and non-glycosylated substrates
J Biol Chem
287, 36473–36487. Rightsholder: AMERICAN SOC FOR BIOCHEMISTRY & MOLECULAR BIOLOGY).
Figure 4.12 Results of the positional scan analysis of library 1344 against ADAM10 and -17. Positional scan of R
1
(a), R
2
(b), R
3
(c), and R
4
(d) defined moieties against ADAM10 (red bars) and ADAM17 (blue bars) using glycosylated substrate. (Adapted from Minond, D., Cudic, M., Bionda, N., Giulianotti, M., Maida, L., Houghten, R. A., and Fields, G. B. (2012) Discovery of novel inhibitors of a disintegrin and metalloprotease 17 (ADAM17) using glycosylated and non-glycosylated substrates.
J Biol Chem
287, 36473–36487. Rightsholder: AMERICAN SOC FOR BIOCHEMISTRY & MOLECULAR BIOLOGY).
Figure 4.13 Results of dose response study of most ADAM17 selective and potent individual compounds. Structures of individual compounds are shown as inserts. (Adapted from Minond, D., Cudic, M., Bionda, N., Giulianotti, M., Maida, L., Houghten, R. A., and Fields, G. B. (2012) Discovery of novel inhibitors of a disintegrin and metalloprotease 17 (ADAM17) using glycosylated and non-glycosylated substrates.
J Biol Chem
287, 36473–36487. Rightsholder: AMERICAN SOC FOR BIOCHEMISTRY & MOLECULAR BIOLOGY).
Figure 4.14 Characterization of mechanism of inhibition of ADAM17 catalytic domain and ectodomain by compound #15. (a) Yonetani-Theorell plot of glycosylated substrate hydrolysis by ADAM17 in the presence of AHA and compound #15. Note the non-parallel lines of best fit indicating mutually non-exclusive binding by two inhibitors. Structure of N-hydroxyacetamide (AHA) shown as insert. (b) Lineweaver-Burke plot of glycosylated substrate hydrolysis by ADAM17 in the presence of compound #15. Dose response study of inhibition of ADAM17 catalytic domain and ectodomain by (c) AHA and (d) compound #15. (Adapted from Minond, D., Cudic, M., Bionda, N., Giulianotti, M., Maida, L., Houghten, R. A., and Fields, G. B. (2012) Discovery of novel inhibitors of a disintegrin and metalloprotease 17 (ADAM17) using glycosylated and non-glycosylated substrates
J Biol Chem
287, 36473–36487. Rightsholder: AMERICAN SOC FOR BIOCHEMISTRY & MOLECULAR BIOLOGY).
Chapter 7: Meprin and ADAM Metalloproteases: Two Sides of the Same Coin?
Figure 7.1 Domain structure and function of meprin α, meprin β and ADAM17. The functions of the domains are indicated in the figure.
Figure 7.2 Physiological functions of ADAM17, meprin α, and meprin β. Both proteases orchestrate different processes in development and during the activation of the immune system.
Chapter 8: Subtracting Matrix out of the Equation: New Key Roles of Matrix Metalloproteinases in Innate Immunity and Disease
Figure 8.1 (a) All 773 reported human MMP substrates distributed for each of the 23 human MMPs. (b) All 773 reported human MMP substrates: the ECM substrates are shown in blue and the non-ECM substrates are shown in green.
Figure 8.2 (a) Gene Ontology (GO) terms enrichment of all 246 reported non-ECM human MMP substrates. (b) Pathway enrichment analysis of the 246 reported non-ECM human MMP substrates.
Chapter 9: MMPs: From Genomics to Degradomics
Figure 9.1 Biological activity of an individual MMP within a local tumor microenvironment. MMPs are central regulators of tumor extracellular environment in terms of both extracellular matrix (ECM) turnover and the signaling milieu controlling cell function. Proteolytic balance is tightly controlled at the protein level by activation of individual MMPs from inactive zymogens (proMMPs) and by the binding of inhibitors. Upon activation, each MMP mediates specific effects on the local microenvironment, dependent on its substrate repertoire. These effects derive either down-stream of the MMPs individual ECM substrates or via activation and/or inactivation of signaling molecules, such as cytokines and growth factors. In consequence, the proteolytic balance influences gene expression and behavior of cancer as well as stromal cells, which in turn are major determinants of the proteolytic balance, the local ECM composition and signaling milieu.
Figure 9.2 Local proteolytic network consisting of interrelated protease systems. The biological effects of an individual MMP (as depicted in Figure 9.1.) are embedded in the interaction with other MMPs that exhibit partially over-lapping but also distinct substrate specificities, forming the local MMP system. Individual proteolytic systems are interrelated, mutually influencing each other in the modulation of protease activity, substrate availability, and action within a tissue.
Figure 9.3 Interconnectivity of local proteolytic networks within an organism. Local proteolytic tissue networks (as depicted in Figure 9.2.) within an organism communicate with each other over a distance via the circulatory system, forming the proteolytic internet. Information is transmitted systemically via up-regulation or down-regulation of soluble factors such as cytokines, hormones, as well as secreted protease inhibitors such as TIMP-1 and PAI-1. The status of homeostasis in the regional proteolytic network of an organ is thereby reported to other tissues in the body. Accordingly, any manipulation of a single member of the proteolytic network results in a re-adaptation. This process is subject to a multitude of net effects that altogether impact on the formation of a new homeostasis, which determines the susceptibility of the organism to disease.
Figure 9.4 Degradomes and degradomics approaches. The transcriptional degradome defines the translational degradome, of which the activity degradome represents the active proteases. Individual active proteases give rise to partial overlapping substrate degradomes. All together, they define the proteolytic potential of a system. For each level of complexity powerful degradomics techniques have been developed. CLIP-CHIP
TM
, Hu/Mu ProtIn, dedicated protease microarrays; SRM, selected reaction monitoring; STEP, STandard of Expressed Protein peptides; ABPs, activity-based probes; PSPs, proteolytic signature peptides; TAILS, terminal amine isotopic labeling of substrates; COFRADIC, combined diagonal fractional chromatography; Subtiligase, engineered peptide ligase for modification of protein N termini.
Figure 9.5 Integrated strategy to elucidate physiological MMP substrates. Multiple candidate substrates from unbiased
in vitro
and cell-based experiments serve as templates for the development of targeted SRM assays that are applied in appropriate
in vivo
models. KO, knockout; WT, wild-type; SRM, selected reaction monitoring.
Chapter 10: MMPs in Biology and Medicine
Figure 10.1 Basic domain structure of MMP and ADAM family members. The characteristic domain structure of MMPs includes (i) the signal peptide domain, which guides the enzyme into the rough endoplasmic reticulum during synthesis, (ii) the propeptide domain, which sustains the latency of these enzymes until it is removed or disrupted, (iii) the catalytic domain, which houses the highly conserved Zn
2+
binding region and is responsible for enzyme activity, (iv) the hemopexin domain, which determines the substrate specificity of MMPs, and (v) a small hinge region, which enables the hemopexin region to present substrate to the active core of the catalytic domain. The subfamily of membrane-type MMPs (MT-MMPs) possesses an additional transmembrane domain and an intracellular domain. MMPs are produced in a latent form and most are activated by extracellular proteolytic cleavage of the propeptide. MT-MMPs also contain a cleavage site for furin proteases, providing the basis for furin-dependent activation of latent MT-MMPs prior to secretion. ADAMs are multidomain proteins composed of propeptide, metalloprotease, disintegrin-like, cysteine-rich, and epidermal growth factor-like domains. Membrane-anchored ADAMs contain a transmembrane and cytoplasmic domain. ADAMTSs have at least one Thrombospondin type I Sequence Repeat (TSR) motif [1]. (Reprinted with permission © (2009) American Society of Clinical Oncology. All rights reserved).
Figure 10.2 Multiple functions of MMPs in cancer progression. (Counterclockwise) MMPs degrade components of ECM, facilitating angiogenesis, tumor cell invasion and metastasis. MMPs modulate the interactions between tumor cells by cleaving E-cadherin, and between tumor cells and ECM by processing integrins, which also enhances the invasiveness of tumor cells. MMPs also process and activate signaling molecules, including growth factors and cytokines, making these factors more accessible to target cells by either liberating them from the ECM (e.g., VEGF and bFGF) and inhibitory complexes (e.g., TGF-β), or by shedding them from cell surface (e.g., HB-EGF) [1]. (Reprinted with permission © (2009) American Society of Clinical Oncology. All rights reserved).
List of Tables
Chapter 2: Dynamics and Mechanism of Substrate Recognition by Matrix Metalloproteases
Table 2.1 Motion parameters of MMPs on substrate surfaces.
Chapter 4: Metzincin Modulators
Table 4.1 Sequences of sarafotoxin analogs.
Table 4.2 Apparent Ki values of Srt variants for different MMPs (μM).
Table 4.3 Amino acid residues encountered in positions 16–20 of TIMPs used to create a combinatorial pool for sarafotoxin S4 engineering.
Table 4.4 Inhibition of MMP-1, MMP-2, MMP-8, MMP-9, MMP-13, and MMP-14 activity by compounds 4, 20, and 24.
Table 4.5 Kinetic parameters for ADAM hydrolysis of glycosylated and non-glycosylated substrates.
Table 4.6 IC
50
values for phage-displayed TIMP-2 variants from the screening of libraries that mutate three regions on TIMP-2.
Chapter 8: Subtracting Matrix out of the Equation: New Key Roles of Matrix Metalloproteinases in Innate Immunity and Disease
Table 8.1 MMP-truncation products of CC chemokines.
Table 8.2 Reported ECM and non-ECM substrates for all 24 human MMPs taken from TopFIND [39].
Table 8.3 Non-proteolytic roles of MMPs.
Chapter 10: MMPs in Biology and Medicine
Table 10.1 Candidate MMP and ADAM biomarkers of cancer.
Table 10.2 Candidate MMP and ADAM biomarkers of non-malignant diseases.
Table 10.3 MMPs/TIMPs as biomarkers for therapeutic efficacy in clinical trials.
Matrix Metalloproteinase Biology
Edited by
Irit Sagi
Professor of Biological Chemistry and Biophysics, Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
Jean P. Gaffney
Assistant Professor of Chemistry at Baruch College, Department of Natural Sciences, City University of New York, New York, NY, USA
Copyright © 2015 by Wiley-Blackwell. All rights reserved
Published by John Wiley & Sons, Inc., Hoboken, New Jersey
Published simultaneously in Canada
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Library of Congress Cataloging-in-Publication Data:
Matrix metalloproteinase biology / edited by Irit Sagi and Jean P. Gaffney.
p. ; cm.
Includes bibliographical references and index.
ISBN 978-1-118-77232-4 (cloth)
I. Sagi, Irit, editor. II. Gaffney, Jean P., editor.
[DNLM: 1. Matrix Metalloproteinases. QU 136]
QP552.M47
572′.696–dc23
2015000037
List of Contributors
Christoph Becker-Pauly
Biochemisches Institut
Medizinische Fakultät
Christian-Albrechts-Universität zu Kiel
Kiel, Germany
Jian Cao
Department of Medicine
State University of New York at Stony Brook
Stony Brook, NY, USA
Jillian Cathcart
Department of Medicine
State University of New York at
Stony Brook Stony Brook, NY, USA
Ivan E. Collier
Departments of Medicine
Division of Dermatology
Washington University School of Medicine
St. Louis, MO, USA
Howard C. Crawford
Department of Cancer Biology
Mayo Clinic
Jacksonville, FL, USA
Antoine Dufour
Department of Oral Biological & Medical
Sciences and Department of Biochemistry and Molecular Biology
Centre for Blood
Research University of British Columbia
Vancouver, BC, Canada
Gregg B. Fields
Torrey Pines Research Institute for
Molecular Studies
Port St. Lucie, FL, USA
Marco Fragai
Magnetic Resonance Center and Department of Chemistry
University of Florence
Florence, Italy
Gregory I. Goldberg
Departments of Medicine
Biochemistry and Molecular Biophysics
Washington University School of Medicine
St. Louis, MO, USA
Barbara Grünwald
Institute for Experimental Oncology and Therapy Research
Klinikum rechts der Isar
Technische Universität München
Munich, Germany
Di Jia
Vascular Biology Program and Department of Surgery
Boston Children's Hospital and Harvard
Medical School
Boston, MA, USA
Ulrich auf dem Keller
Department of Biology
Institute of Molecular Health Sciences
ETH Zurich,
Zurich, Switzerland
Achim Krüger
Institute for Experimental Oncology and Therapy Research
Klinikum rechts der Isar
Technische Universität München
Munich, Germany
Claudio Luchinat
Magnetic Resonance Center and Department of Chemistry
University of Florence
Florence, Italy
Dmitriy Minond
Cancer Research
Torrey Pines Research Institute for Molecular Studies
Port St. Lucie, FL, USA
Marsha A. Moses
Vascular Biology Program and Department of Surgery
Boston Children's Hospital and Harvard
Medical School
Boston, MA, USA
Christopher M. Overall
Department of Oral Biological & Medical
Sciences and Department of Biochemistry and Molecular Biology
Centre for Blood Research
University of British Columbia
Vancouver, BC, Canada
Ashleigh Pulkoski-Gross
Department of Medicine
State University of New York at Stony Brook
Stony Brook, NY, USA
Stefan Rose-John
Biochemisches Institut
Medizinische Fakultät
Christian-Albrechts-Universität zu Kiel
Kiel, Germany
Roopali Roy
Vascular Biology Program and Department of Surgery
Boston Children's Hospital and Harvard
Medical School
Boston, MA, USA
Pascal Schlage
Department of Biology
Institute of Molecular Health Sciences
ETH Zurich, Zurich, Switzerland
M. Sharon Stack
Harper Cancer Research Institute
University of Notre Dame
South Bend, IN, USA
Maciej J. Stawikowski
Torrey Pines Research Institute
Torrey Pines, FL, USA
Stanley Zucker
VA Medical Center
Northport, NY, USA
Chapter 1Matrix Metalloproteinases: From Structure to Function
Maciej J. Stawikowski1 and Gregg B. Fields2
Departments of Chemistry and Biology, Torrey Pines Institute for Molecular Studies, Port St. Lucie, USA
1.1 Introduction
Members of the matrix metalloproteinase (MMP) family are known to catalyze the hydrolysis of a great variety of biological macromolecules. Proteomic approaches have significantly expanded the number of known MMP substrates. However, the mechanisms by which macromolecular substrates are processed have often proved elusive. X-ray crystallography and NMR spectroscopy have yielded detailed information on structures of MMP domains and, in a few cases, full-length MMPs. As structures of MMPs and their substrates have been reported, examination of MMP•substrate complexes has provided insight into mechanisms of action. We examine the structures of MMPs and their substrates and consider how the various structural elements of MMPs contribute to the hydrolysis of biological macromolecules.
1.2 Structures of MMPs
1.2.1 General MMP structure and domain organization
MMPs belong to the M10 zinc metalloproteinase family [1]. All MMPs have the characteristic zinc binding motif HExxHxxGxxH in their catalytic domain. MMPs possess similar domain organizations. Most MMPs consist of a signal peptide followed by four distinct domains, the N-terminal prodomain (propeptide), catalytic (CAT) domain, linker (hinge) region, and C-terminal hemopexin-like (HPX) domain (Fig. 1.1). The membrane-type (MT) MMPs contain an additional transmembrane (TM) domain that anchors them to the cell membrane. Following the TM domain is a small cytoplasmic “tail”.
Figure 1.1 General domain organization of MMPs.
There are several exceptions to this general domain organization. MMP-7 and MMP-26 (matrilysins) lack the linker region and HPX domain and thus are referred to as “minimal MMPs”. MMP-2 and MMP-9 possess three repeats of fibronectin type II-like motifs within the CAT domain. MMP-17 and MMP-25 are type I TM enzymes anchored to membranes through a C-terminal glycosylphosphatidylinositol (GPI) residue [2]. The N-terminal MMP-23 pro-domain contains a type II TM domain that anchors the protein to the plasma membrane. Instead of the C-terminal HPX domain common to other MMPs, MMP-23 contains a small toxin-like domain (TxD) and an immunoglobulin-like cell adhesion molecule (IgCAM) domain.
1.2.2 Catalytic domain
The topology of the CAT domain is similar among all MMPs. The CAT domain is composed of a five- stranded β-sheet which is interrupted by three α-helices (Fig. 1.2). Four of the five β-strands are aligned in a parallel fashion, while only the smallest “edge” strand runs in the opposite direction. Between strands III and IV there is an S-loop fixed by a structural Zn atom. The center of the catalytic site is located at helix B and the loop connecting it with helix C. This center helix provides the first and second His residues of the Zn-binding motif along with “catalytic” Glu residue. The loop behind this helix provides the third zinc binding His residue. Further down along this loop there is a 1,4 β-turn forming Met residue. This residue is highly conserved among metzincins and is believed essential for the structural integrity of the zinc-binding site. However, MMP-2 mutants where the conserved Met was replaced with Leu or Ser were able to cleave gelatin, type I collagen, and chemokine monocyte chemoattractant protein-3 with similar efficiency as wild-type MMP-2 [3].
Figure 1.2 Typical structure of the CAT domain of MMPs. Characteristic structural elements are highlighted with arrows. Figure generated using MMP-8 structure (PDB 2OY2) [4].
1.2.3 Catalytic mechanism
On the basis of early structural information, a catalytic mechanism for MMPs was proposed (Fig. 1.3) [5, 6]. The carbonyl group of the scissile bond coordinates to the active site zinc (II) ion. A water molecule is hydrogen bonded to a conserved Glu residue and coordinated to the zinc (II) ion. The water molecule donates a proton to the Glu residue, allowing the generated hydroxide ion to attack the carbonyl at the scissile bond. This attack results in a tetrahedral intermediate, which is stabilized by the zinc (II) ion. The Glu residue transfers a proton to the nitrogen of the scissile amide, the tetrahedral intermediate rearranges, and amide bond hydrolysis occurs. During this catalytic process, the carbonyl from a conserved Ala residue helps to stabilize the positive charge at the nitrogen of the scissile amide.
Figure 1.3 Mechanism of proteolysis catalyzed by MMPs. (Figure prepared based on mechanism proposed by Lovejoy et al. [5]).
1.2.4 Fibronectin type II-like inserts
Gelatinases (MMP-2 and MMP-9) bind to gelatin and collagen with significant contribution from their three fibronectin type II-like (FN2) repeats. MMP-2 and MMP-9 are unique among the MMPs in that the three FN2 modules (Col-1, Col-2, and Col-3) are inserted in their CAT domain in the vicinity of the active site [7]. More specifically, the FN2 modules of MMP-2 and MMP-9 are inserted between the fifth β-strand and helix B in the CAT domain (according to active enzyme domain organization). The basic fold of the FN2 module comprises a pair of β-sheets, each made from two antiparallel strands, connected by a short α-helix (Fig. 1.4). The two β-sheets form a hydrophobic pocket that is part of a hairpin turn, which orients the surrounding aromatic side chains into the hydrophobic pocket. These pockets are the structural hallmark of the FN2 modules and contribute to substrate binding (see below) [8].
Figure 1.4 Fibronectin type II-like module structure and organization. (a) General orientation of FN2 modules of MMP-2. (b) Top view of FN2 modules. Figure prepared using MMP-2 structure (PDB 1CK7) [8].
1.2.5 Linker region
The CAT domain is connected to the HPX domain via a linker (hinge) region. The length of this linker varies from 8 to 72 amino acids, depending on the enzyme (Fig. 1.5). The linker regions may be posttranslationally modified with sugar moieties. The conformational flexibility of the linker region contributes to MMP function. For example, in the case of MMP-9, it has been suggested that the long (72 residue), glycosylated, and flexible linker region mediates protein-substrate interactions by allowing the independent movement of the enzyme CAT and HPX domains [9]. Independent domain movements were also proposed to mediate enzyme translocation on collagen fibrils [10–12]. Domain flexibility may contribute to MMP activation via promoting long-range conformational transitions induced by the binding of activator proteins or ligand [13–15]. Finally, the linker region may help to re-orient the CAT domain with respect to the HPX domain during catalysis of collagen [16]. Domain flexibility may be rationalized for most MMPs by considering the amino acid composition (i.e., Gly and Pro residues) and the various lengths of linker regions (Fig. 1.5). The linker region and HPX domain of MT1-MMP and MMP-9 are proposed to offer allosteric control of enzyme dimer formation, which in turn modulates biological function [17, 18].
Figure 1.5 Comparison of MMP linker lengths and sequences. Table was generated after alignment of human MMPs using sequences from the Uniprot database [19] and SeaView 4 [20] and Jalview [21] programs.
Glycosylation of MT1-MMP, which occurs in the linker region (residues 291, 299, 300, and 301), is required for the recruitment of tissue inhibitor of metalloproteinase 2 (TIMP-2) on the cell surface and subsequent formation of the MT1-MMP/TIMP-2/proMMP-2 trimeric complex and activation of proMMP-2 [22]. Glycosylation does not affect MT1-MMP collagen hydrolysis or autolytic processing [22].
1.2.6 Hemopexin-like domain
Except for MMP-7 and MMP-26, all vertebrate and human MMPs are expressed with a C-terminal HPX domain. The HPX domain is organized in four β-sheets (I to IV), arranged almost symmetrically around a central axis in a consecutive order (Fig. 1.6). The end result is a four-bladed propeller of pseudo-fourfold symmetry. Each propeller blade is formed by four antiparallel β-strands connected in a W-like topology, and is strongly twisted. The small C-terminal helix of the blade IV is tethered to the entering strand of blade I via a single disulfide bridge, stabilizing the whole domain. Within the central tunnel, up to four ions (2Ca2+, 2Cl−) have been identified although their function is not clear [23].
Figure 1.6 Typical structure of the HPX domain. The propeller-like structure is composed of four blades (I-IV) and stabilized by a single disulfide bridge, designated with an arrow. In the central tunnel, up to four different ions have been identified (here Ca2+ is orange and Cl− is yellow). This figure was generated using the HPX domain of MT1-MMP (PDB 3C7X) [23].
The HPX domain mediates binding of MMP-1, MMP-8, MMP-13, MT1-MMP, and MMP-3 to collagen [24–28]. The HPX domain of MMP-2 was shown to possess critical secondary binding sites (exosites) required for the interactions of MMP-2 with fibronectin, and fibronectin was cleaved at a significantly reduced rate by an MMP-2 variant where the HPX domain was deleted [29]. In the case of MMP-2 and MMP-9, the HPX domain is important for interactions with TIMPs. The HPX domain of MMP-2 has also been shown to play a role in zymogen activation by MT1-MMP [30].
HPX domains modulate interaction of MMPs with cell-surface biomolecules. For example, the HPX of MMP-2 plays a role in the binding of the enzyme to the αvβ3 integrin [31, 32]. MT1-MMP has numerous cell surface binding partners, including tetraspanins (CD9, CD63, CD81, CD151, and/or TSPAN12), the α2β1 and αvβ3 integrins, and CD44 [33–39]. The HPX domain of MT1-MMP binds to CD63 and CD151 [35, 40]. Tetraspanins protect newly synthesized MT1-MMP from lysosomal degradation and support delivery to the cell surface [36].
CD44 also binds to MT1-MMP via the HPX domain of the enzyme, specifically blade I of the HPX domain [34, 41]. The association with CD44 leads to MT1-MMP localization to lamellipodia [34, 40]. The MT1-MMP/CD44 interaction promotes signaling through EGFR activation to the MAPK and PI3K pathways, enhancing cell migration [41]. CD44 also binds to MMP-9 via the HPX domain [40].
Highly efficient collagenolysis requires homodimerization of MT1-MMP, where association includes interactions of the HPX domain [42]. Homodimerization is symmetrical, involving residues Asp385, Lys386, Thr412, and Tyr436 in blades II and III of the HPX domain [43].
1.2.7 Transmembrane domain and cytoplasmic tail
On the basis of their method of attachment to the cell membrane, MT-MMPs may be classified into two groups, TM-type and glycosylphosphatidyl-inositol (GPI)-type. MT1-MMP (MMP-14), MT2-MMP (MMP-15), MT3-MMP (MMP-16), and MT5-MMP (MMP-24) are type I TM proteins with a short cytoplasmic tail that is involved in the regulation of intracellular trafficking and activity of these proteases [44–46]. MT4-MMP (MMP-17) and MT6-MMP (MMP-25) are bound to the cell surface by a GPI-mediated mechanism [2, 47].
Although the structure of the TM domain has not been solved experimentally, a model has been generated (Fig. 1.7). Besides facilitating cellular localization, the TM domain allows MT-MMPs to process a unique set of substrates, interact uniquely with TIMPs, and participate in a non-conventional mechanism of regulation involving enzyme internalization, processing, and ectodomain shedding [48, 49].
Figure 1.7 Structure of TM domain and cytoplasmic tail (residues 518–582) of human MT1-MMP generated by homology modeling [50, 51].
The cytoplasmic tail of MT1-MMP is distinct from those of MT2-MMP, MT3-MMP, and MT5-MMP, and is well characterized. The cytoplasmic tail of MT1-MMP is important in the ERK activation cascade [52], S1P-dependent Gi protein signaling [53], and VEGF upregulation through Src tyrosine kinase pathways [54]. The multifunctional gC1qR proteins can bind to the cytoplasmic tail of MT1-MMP in a similar manner to the cytoplasmic portion of adrenergic receptor [55]. More recently, Uekita et al. [56] have identified a new 19 kDa MT1-MMP cytoplasmic tail binding protein-1 (MTCBP-1). MTCBP-1 is localized between three subcellular compartments (membrane, cytoplasm, and nucleus) that can regulate gene expression and may suppress the invasion and migration-promoting activity of MT1-MMP [56]. The cytoplasmic tail of MT1-MMP increases the expression of hypoxia inducible factor-1 (HIF-1) target genes, which in turn stimulates aerobic glycolysis [57].
Phosphorylation of cytoplasmic Tyr573 of MT1-MMP is imperative for tumor cell migration and proliferation in three-dimensional collagen matrices and tumor growth in mice [58, 59], while phosphorylation of cytoplasmic Thr567 enhances tumor invasion of and growth within collagen matrices [60]. Interestingly, Tyr573 phosphorylation does not affect proteolytic activity, but may act by inducing relocalization of the enzyme and increasing the proportion of quiescent cells [58].
MT1-MMP undergoes both clathrin-mediated and caveolae-dependent endocytosis [61, 62, 42]. The cytoplasmic tail has been implicated as necessary for endocytosis [61].
1.3 Overview of MMP substrate specificity
Extensive sequence specificity studies of many MMPs provided a number of important insights into the differences and similarities in subsite preferences among these enzymes. Substrate specificity studies have been performed with proteins and synthetic peptides.
Significant interactions between the MMPs and their substrates or inhibitors occur between the S1′ subsite and P1′ residue. MMPs may be classified as falling into two broad structural classes dependent on the depth of the S1′ pocket. This “selectivity pocket” is relatively deep for the majority of the enzymes (e.g., MMP-2, MMP-3, MMP-8, MMP-12, and MMP-13) but shallow in the case of MMP-1, MMP-7, and MMP-11.
The substrate-binding groove is relatively open at the S3-S1 and S3′ subsites and narrows at the S1′ and S2′ subsites. The S1′ subsite is a well-defined pocket that penetrates the surface of the enzyme. Differences between the various MMPs in the S3–S1 subsite region are relatively subtle. Interestingly, Pro is a preferred P3 subsite moiety for many MMP substrates.
The S2′ subsite is a solvent-exposed cleft with a general preference for hydrophobic P2′ residues in both substrates and inhibitors. The S3′ subsite is a relatively poorly defined solvent-exposed region. While there are some variations in residues for this subsite for the various MMPs, the introduction of different P3′ substituents in general tends to have only a modest effect on inhibitor selectivity.
In addition to active site subsites, the specificity of MMPs is modulated by discrete binding sites outside of the catalytic center (exosites). Substrate interaction with exosites can influence the behavior of a proteinase in a number of ways. Exosites modulate and broaden the substrate specificity profile of MMPs by providing an additional contact area not influenced by the primary specificity subsites. In this way, the function of the proteinase is refined and can be made, in general, more specific or efficient. In addition to bringing substrates to the enzyme for potential hydrolysis, exosites may be involved in essential “substrate preparation” prior to cleavage. For example, the localized “unwinding” of native collagen substrates by MMPs is facilitated by exosites [16]. Exosites can also target the enzyme to substrates in tissues or to cell-associated substrates.
In collagenolytic MMPs (MMP-1, MMP-8, MMP-13, and MT1-MMP), exosites are found in the HPX domain, and in gelatinases (MMP-2 and MMP-9), on the three FN2 modules. In MMP-1, MMP-8, MMP-13, MT1-MMP, and MMP-3, the HPX domain binds native collagen. The FN2 modules in gelatinases form a collagen binding domain (CBD) which lies proximal to the S3′ subsite. The matrix binding properties of the CBD also have the potential to localize the enzyme to collagen, either in the extracellular matrix (ECM) or on the cell surface linked to β1 integrins.
1.3.1 ECM substrates
The repertoire of MMP substrates is extremely rich. To study proteolytic processes in detail (referred to as the protease web), a broad approach including gene deletion, transgenic mouse models, and genomic and proteomic profiling techniques is necessary. Degradomics, the characterization of all proteases, inhibitors, and protease substrates present in an organism using genomic and proteomic techniques, is a well-established method for MMP substrate identification [63].
The ECM is composed of two main classes of macromolecules: proteoglycans (PGs) and fibrous proteins [64, 65]. The main fibrous ECM proteins are collagens, elastin, fibronectin, and laminins [66]. PGs fill the majority of the extracellular interstitial space within the tissue in the form of a hydrated gel [64].
Collagen is the most abundant fibrous protein within the interstitial ECM and constitutes up to 30% of the total protein mass of a multicellular animal. Collagens provide tensile strength, regulate cell adhesion, support chemotaxis and migration, and direct tissue development [67]. Collagen associates with elastin, another major ECM fiber. Elastin fibers provide recoil to tissues that undergo repeated stretch. A third fibrous protein, fibronectin, is intimately involved in directing the organization of the interstitial ECM and has a crucial role in mediating cell attachment and function.
Collagenases (MMP-1, MMP-8, MMP-13, and MT1-MMP) catalyze the degradation of fibrillar collagens in their native triple-helical supersecondary structure. The physiological role of collagenases has been proposed to be the remodeling of the collagenous component of the ECM, including involvement in the wound healing process. Furthermore, since collagen is the predominant ECM deposit in fibrotic organs, collagenases are believed to be the main proteases responsible for the resolution of fibrosis and restoration of the normal ECM environment. Numerous ECM components, including types I, II, and III collagen, fibronectin, vitronectin, laminins 111 and 332, fibrin, and proteoglycans are substrates for MT1-MMP [68].
Gelatinases (MMP-2 and MMP-9) have been proposed to be involved in inflammatory processes and in tumor progression [69, 70]. However, gelatinases have also been found to have protective roles against cancer [71–74]. Gelatinases have been more recently recognized as participating in cardiovascular and auto-immune diseases. In the case of cardiovascular diseases, gelatinases participate in both the genesis of atherosclerotic lesions and to the acute event (i.e., stroke or myocardial infarction). In the case of auto-immune diseases, gelatinases are involved in the generation of remnant epitopes and in the modulation of cross-talk between immune system compartments.
Stromelysins (MMP-3, MMP-10, and MMP-11) share the ability to degrade types IV and IX collagen, laminin, fibronectin, elastin, and proteoglycans, although with significantly different affinities among them. Additional substrates include cytokines, growth factors, and soluble regulatory molecules [75]. Each stromelysin has a different physiological distribution in human tissues, hence the types of processes which are modulated are largely variable.
Among the matrilysins (MMP-7 and MMP-26), MMP-7 is widely expressed in human tissues and mainly in epithelial-derived ones. MMP-7 catalyzes the hydrolysis of cytokines, growth factors, and receptors [76]. MMP-7 biological functions mainly concern ECM remodeling and immune system modulation. The biological aspects of MMP-26 are so far restricted to ECM turnover and remodeling in a limited cohort of tissues both in physiological and pathological conditions [77, 78].
The matrix metalloproteinase term initially related to enzymes processing ECM proteins, but recent findings prove that the role of MMPs is much more sophisticated. MMPs contribute to processing of cytokines, chemokines, hormones, adhesion molecules, and membrane-bound proteins, resulting in modulation of normal cellular behavior, cell-cell communication, and tumor progression. The reader is referred to several excellent reviews on MMPs that have compiled an extensive list of substrates [79–81].
1.3.2 Cell surface substrates
Proteolytic events at the cell surface are of interest because of their potential to affect cellular functions. Cell surface-associated MMP-2, MMP-9, and MMP-13 can activate latent transforming growth factor-beta (TGFβ) [82]. MT1-MMP modulates the bioavailability of TGFβ (i) by activating MMP-13 and MMP-2 [83], (ii) by releasing active TGFβ from cell surface complexes involving the αvβ3 integrin [84], and/or (iii) by releasing a membrane-anchored proteoglycan, betaglycan, that binds TGFβ [85].
MT-MMPs can cleave and shed a variety of cell surface adhesion receptors and proteoglycans. CD44 (a multifunctional adhesion molecule) [86] and syndecan-1 [87] can be directly shed by MT1-MMP and MT3-MMP. The αv chain of the αvβ3 integrin, which is reported to play a crucial role in tumor angiogenesis, invasion, and metastasis, is processed by MT1-MMP into a functional form [88]. The multifunctional receptor of complement component 1q (gC1qr) is also susceptible to MT1-MMP proteolysis [89]. Low-density lipoprotein receptor related protein (LRP1/CD91) is a cell surface-associated endocytic receptor, implicated in the internalization and degradation of multiple ligands such as thrombospondins (1 and 2), α2-macroglobulin-protease complexes, urokinase- and tissue-type plasminogen activators, MMP-2, MMP-9, and MMP-13 [90, 91]. The cleavage of LRP1 by MT1-MMP in breast cancer and fibrosarcoma cells may thus lead to the control of the bioavailability and fate of many ligands and soluble MMPs in cancer progression [91]. MT1-MMP also sheds transglutaminase (Belkin et al., 2001), death receptor-6, MHC class I chain-related molecule A, E-cadherin, and ECM metalloproteinase inducer [92–96]. These highly divergent substrates for MT1-MMP make this enzyme a critical regulator of the pericellular environment.
1.3.3 Intracellular MMP targets
For a long time MMPs were viewed exclusively as ECM remodelers. More recently, there is evidence that MMPs cleave intracellular substrates, and that MMPs have been observed within cells in nuclear, mitochondrial, and various vesicular and cytoplasmic compartments, including the cytoskeletal intracellular matrix. Unbiased high-throughput degradomics approaches have demonstrated that many intracellular proteins are cleaved by MMPs, including apoptotic regulators, signal transducers, molecular chaperones, cytoskeletal proteins, systemic autoantigens, enzymes in carbohydrate metabolism and protein biosynthesis, transcriptional and translational regulators, and proteins in charge of protein clearance such as lysosomal and ubiquitination enzymes. Intracellular substrate proteolysis by MMPs is involved in innate immune defense and apoptosis, and affects oncogenesis and pathology of cardiac, neurological, protein conformational, and autoimmune diseases, including ischemia-reperfusion injury, cardiomyopathy, Parkinson's disease, cataract, multiple sclerosis, and systemic lupus erythematosus. Intracellular activation of MMPs strongly suggests that MMPs are responsible for proteolytic actions on intracellular substrates.
MMP-2 cleaves the cytoskeletal proteins desmin and α-actinin and colocalizes with α-actinin in cardiomyocytes [97]. MMP-2 and MMP-9-containing vesicles are aligned with the cytoskeleton in neurons and reactive astrocytes, and both gelatinases are found in cytoskeletal fractions from these cells [98, 99]. MT1-MMP and MT3-MMP are detected in cytoskeletal fractions of smooth muscle cells, where they cleave the cytoskeletal protein focal adhesion kinase (FAK) [100]. Moreover, cytoskeletal proteins constitute an important fraction of the intracellular degradomes of MMP-2, MMP-9, and MT1-MMP. Both pro- and activated MMP-1 are associated with the mitochondrial membrane in glial Müller cells, Tenon's capsule fibroblasts, corneal fibroblasts, and retinal pigment epithelial cells [101]. The mitochondrial localization of MMP-1 is found in resting cells, suggesting a physiological role for MMP-1 in cellular homeostasis. Both MMP-2 [102, 103] and MMP-9 [104] are detected in cardiac mitochondria during cardiac injury and increased levels of mitochondrial MMP-9 are associated with exacerbated mechanical dysfunction. Studies report nuclear localization of MMPs, including MMP-1, MMP-2, MMP-3, MMP-9, MMP-13, MMP-26, and MT1-MMP, and cleavage of nuclear matrix proteins. Nuclear translocation of MMP-3 was confirmed by Eguchi et al. [105], who showed that extracellular MMP-3 is taken up into chondrosarcoma cells and subsequently translocates to the nucleus where it induces transcription of the connective tissue growth factor (CTGF) gene. To avoid excessive proteolysis of nuclear proteins during cellular homeostasis, these nuclear MMPs may be under inhibition by TIMP-1 and TIMP-4, which are also present in the nucleus [106–109]. MMP-7 colocalizes with cryptdins (antimicrobial α-defensins, Crps) in mouse Paneth cells and mediates the processing and activation of various Crps in vitro [110]. MMP-7 cleaves pro-Crp-1, −6, and −15. MT1-MMP was shown to have an intracellular oncogenic function by cleaving the integral centrosomal protein, pericentrin [111, 112]. Pericentrin and pericentrin-2 (pericentrin-B or kendrin) are derived from splice variants of the same gene and are known to be essential for normal centrosome function by the anchorage of the γ-tubulin ring complex, which initiates microtubule nucleation, to the centrosome [113]. Besides its actions in the centrosomal compartment and at focal adhesions, activated MT1-MMP is also detected in the nuclei of hepatocellular carcinoma cells. Interestingly, liver cancer patients with nuclear MT1-MMP (and co-localized MMP-2) have a poor overall survival and large tumor size, whereas MT1-MMP is not found in nuclei of normal paralleled liver tissues and normal control livers [114]. Finally, MMP-1 was found to be strongly associated with mitochondrial membranes and nuclei and accumulated within cells during the mitotic phase of the cell cycle. The intracellular association of MMP-1 to mitochondria and nuclei conferred resistance to apoptosis, which may be a mechanism for tumor cells to escape from apoptosis [101].
MMP-2 was localized to sarcomeres in close association with the thin myofilaments in hearts subjected to ischemia/reperfusion (I/R) [102]. Interestingly, a different localization of the two gelatinases, MMP-2 and MMP-9, was observed in hearts of patients with dilated cardiomyopathy (DCM) compared to control hearts. In DCM hearts the gelatinases were localized exclusively within the cardiomyocytes in close association with the sarcomeric structure, whereas localization was mainly around the myocytes in control hearts. I/R injury is associated with the degradation of cytoskeletal proteins such as α-actinin, desmin, and spectrin [115]. This may constitute an additional intracellular function of MMP-2, as α-actinin and desmin (but not spectrin) were found to be in vitro substrates of MMP-2. Moreover, dopaminergic neuroblastoma cells under oxidative stress showed an upregulation of intracellular and secreted3 activated forms of MMP-3 and cleavage of α-synuclein, which was inhibited by an MMP inhibitor. Purified α-synuclein is cleaved by MMP-3 most efficiently, but also by MT1-MMP, MMP-2, MMP-1, and MMP-9 (ordered by decreasing efficiency) [116].
1.4 Selective mechanisms of action
1.4.1 Collagenolysis
Collagens are composed of three α chains of primarily repeating Gly-Xxx-Yyy triplets, which induce each α chain to adopt a left-handed polyPro II helix. Three chains then intertwine, staggered by one residue and coiled, to form a right-handed superhelix [117, 118]. Triple-helical structure provides collagens with exceptional mechanical strength and broad resistance to proteolytic enzymes. Interstitial collagens have long been recognized as being hydrolyzed by collagenolytic MMPs (MMP-1, MMP-8, MMP-13, and MT1-MMP) into one-fourth and three-fourth length fragments.
The 15 Å collagen triple-helix does not fit into the 5 Å MMP CAT domain active site cavity [119]. Models have generally accounted for the steric clash of the triple-helix with enzyme-active sites by (i) requiring active unwinding of the triple-helix by an MMP [119–121] and/or (ii) considering that the site of hydrolysis within collagen has a distinct conformation, or conformational flexibility, rendering it more susceptible to proteolysis than other regions in collagen [122].
A detailed mechanism of collagenolysis was developed from examination of structures and docking experiments of MMP-1 and MMP-1•triple-helical peptide (THP) complexes [16]. MMP-1 in solution is in equilibrium between open/extended and closed structures (Fig. 1.8(a)) [12]. The maximum occurrence (MO) of MMP-1 conformations in solution has recently been calculated, through paramagnetic NMR and small angle X-ray scattering [123]. Many of the MMP-1 conformations with the highest MO value (>35%) were found to have interdomain orientations and positions that could be grouped into a cluster [123]. Within this cluster, the collagen- binding residues of the HPX domain were solvent-exposed and the CAT domain correctly positioned for its subsequent interaction with the collagen. A approximately 50° rotation around a single axis of the CAT domain with respect to the HPX domain positioned the CAT domain right in front of the preferred cleavage site in interstitial collagen. The conformations belonging to this cluster can thus be seen as the antecedent step of collagenolysis.
Figure 1.8 Mechanism of the initial steps of collagenolysis. (a) Closed (left) and open/extended (right) forms of MMP-1 in equilibrium. (b) The extended protein binds THP chains 1T-2T at Val23-Leu26 with the HPX domain and the residues around the cleavage site with the CAT domain. The THP is still in a compact conformation. (c) Closed FL-MMP-1 interacting with the released 1T chain (in magenta). (d) After hydrolysis, both peptide fragments (C- and N-terminal) are initially bound to the active site. (e) The C-terminal region of the N-terminal peptide fragment is released. (Reprinted with permission from [16]. Copyright (2012) American Chemical Society).
The HPX domain then binds the leading chain (designated 1T) and the middle chain (designated 2T) of the THP and, due to the flexibility of the linker, the CAT domain is guided toward the Gly˜Ile bond of chain 1T (Fig. 1.8(b)). Back-rotation of the CAT and HPX domains, to achieve the X-ray crystallographic closed MMP-1 conformation, resulted in visible perturbation of the THP (Fig. 1.8
