ARTICLE pubs.acs.org/Langmuir
Investigation of Early Stages of Fibrin Association Elena G. Zavyalova,*,† Anna D. Protopopova,‡,§ Aleksey M. Kopylov,† and Igor V. Yaminsky†,‡,§ †
Chemistry Department and ‡Physical Department, M. V. Lomonosov Moscow State University, Leninskie gory 1-3, Moscow, 119991 Russian Federation § Advanced Technologies Center, Stroitelei street 4-5-47, Moscow, 119311 Russian Federation ABSTRACT: Interactions between fibrinogen molecules proteolytically cleaved with thrombin were investigated using atomic force microscopy (AFM) and dynamic light scattering (DLS). Gradually decreased fibrinogen concentrations were used to study the fibrin network, large separated fibrils, small fibrils in the initial association stages, and protofibrils. In addition, a new type of structure was found in AFM experiments at a low fibrinogen concentration (20 nM): the molecules in these single-stranded associates are arranged in a row, one after the other. The height, diameter, and distance between domains in these single-stranded associates were the same as those in the original fibrinogen molecules. DLS data assumed formation of extended associates in bulk solution at fibrinogen concentration as low as 20 nM.
’ INTRODUCTION Fibrinogen and thrombin are key proteins in blood clotting. Fibrinogen is a 340 kDa fibrous glycoprotein consisting of three pairs of nonidentical polypeptide chains (R, 610 residues; β, 461 residues; γ, 411 residues). It is 45 nm long and 25 nm wide. The N-terminal sequences are coupled with disulfide bonds into the E domain in the center of the molecule. The C-terminal sequences of nonidentical chains are held together into the D domain, and fibrinogen hence has two D domains on the ends. The region between the E and the D domains is mainly a threestranded R-helical coiled-coil structure stabilized by disulfide bonds. This particular feature gives fibrinogen considerable flexibility.1,2 Thrombin is a highly specific serine protease that cleaves only 4 out of 376 tripsin-sensitive fibrinogen peptide bonds. These bonds are located in the N-terminal sequences of R and β chains (E domain), and four fibrinopeptides are therefore cleaved: two fibrinopeptides A with 16 amino acids each and two fibrinopeptides B with 14 amino acids each.1,2 Cleavage of all fibrinopeptides yields fibrin, and the structure and mass of fibrin are similar to the original fibrinogen. Fibrin spontaneously associates to form fibers a few micrometers long and up to 200 nm wide. In accordance with the widely accepted fiber formation model, a key association stage involves newly generated N-terminal sequences (in the E domain) interacting with specific sites in the D domains. As a result, each E domain is coupled to two D domains, forming double-stranded associates (protofibrils) that in turn join into thicker fibers.2,3 Protofibrils and mature fibrin fibers were previously investigated by electron microscopy47 and atomic force microscopy (AFM).810 Fibrinogen adsorption on different substrates has been studied.1116 O’Brien et al. described an alternative mechanism of fibrin association. Extensive flexible molecularly thin two-dimensional r 2011 American Chemical Society
sheets are formed at low fibrinogen and thrombin concentrations (about 60 and 0.9 nM respectively), and they tend to fold and roll up into thicker fibers. O’Brien et al. supposed that the described sheets could be very short-lived at high fibrinogen concentrations and were therefore missed in earlier studies. The significance of such a fiber formation mechanism in vivo is unclear.17 Koo et al. observed fibrinogen fiber formation at physiological protein concentrations (approximately 10 μM) under nonenzymatic conditions by adsorption on a highly hydrophobic surface (trioctylmethylamine grafted onto silica clay plates). The dimensions and cross-sectional banding period of the observed fibers were close to those of thrombin-induced fibers. Treatment with an RC domain (residues AR 221610) or RC domain antibody significantly reduced the fiber width under such conditions.9 Litvinov et al. determined that the RC domain has an affinity for the N-terminal β-chain fibrinogen sequence but not for the thrombin-cleaved sequence.18 Therefore, at least the RC domain, in particular, in fibrinogen is probably associated with the β-chain sequence and is released after fibrinopeptide B proteolysis with thrombin. This process could be a basis for fiber thickening. However, it is unclear whether the RC domain participates in forming early fibrin associates. This paper presents an investigation of the initial fibrin associate in order to improve our understanding of the fiber formation mechanism. AFM was used as a powerful tool for investigating molecular interactions. The DLS method was used to prove that AFM images reflect fibrin association in a bulk solution. Received: January 13, 2011 Revised: March 3, 2011 Published: March 23, 2011 4922
dx.doi.org/10.1021/la200148n | Langmuir 2011, 27, 4922–4927
Langmuir
ARTICLE
Figure 1. Height mode AFM images of fibrinogen molecules on mica (A, B), on HMDS-mica (C), and on HOPG (D). The color palette has a Z range of 5 nm in A and B, 1.5 nm in C, and 7 nm in D. (B) Zoomed images of fibrinogen molecules on mica, and the schemes below the images represent the possible location of fibrinogen domains on the surface (fibrinogen E domains are shown in black and D domains in yellow).
’ MATERIALS AND METHODS Chemicals. Inorganic salts (analytical grade) were purchased from MP Biomedicals (USA). Recombinant human thrombin with a specific activity of 3.6 kIU/mg (HTI, USA), human thrombin with a specific activity of 110 IU/ml (NIBSC, U.K.), and fibrinogen from human plasma (Calbiochem, Germany) were used. All reactions were carried out at 27 °C in a buffer containing 20 mM Tris-acetate, pH 7.4, 0.14 M NaCl, 50 mM KCl, 10 mM MgCl2, and 10 mM CaCl2. Hexamethyldisilazane (HMDS) from Sigma-Aldrich (USA) was used to modify the mica for AFM measurements. Thrombin Activity Determination. Thrombin activity determination was used to control protease quality during storage and freezingthawing cycles. A 70 μL aliquot of 2 μM fibrinogen solution in buffer was placed into a microcuvette (Uvette; Eppendorf AG, Germany) for the spectrophotometer (Biophotometer; Eppendorf AG, Germany). Typically, 1 μL of thrombin (final concentration 3.6 nM) was added to initiate fibrin network formation. Absorption at 320 nm was measured every 7 s, and the slope of the turbidity curve was determined. Specific thrombin activity in the probe was calculated using standard thrombin activity data. Dynamic Light Scattering (DLS) Experiments. Typically, 300 μL of 2 μM, 0.2 μM, or 20 nM fibrinogen solution was placed into a polystyrene semimicrocuvette (Greiner Bio-One, Germany) and 4 μL of thrombin (final concentration 3.6 nM, specific activity 1.4 kIU/mg) was added to initiate fibrin network formation. Measurements were performed using a Zetasizer Nano (Malvern Instruments, U.K.). All experiments were conducted at least three times. The data were analyzed using Zetasizer Software (Malvern Instruments, U.K.). AFM Measurements. The AFM experiments were conducted using a Multimode AFM with a Nanoscope IIIA Controller (Digital
Instruments, USA) in tapping mode with a typical scan rate of 1 Hz. The measurements were performed in air in tapping mode using standard silicon cantilevers (NT-MDT, Russia) with a guaranteed tip radius of 25 nm and sharp silicon cantilevers (NT-MDT, Russia) with a guaranteed tip radius of 10 nm. For control experiments the protein solution was allowed to adsorb on the surface for a certain time required in each experiment (the times are shown in the figure captions), the solution was then carefully removed with filter paper, the substrate was immediately placed onto the drop of double-distilled Millipore water (this procedure was repeated twice), and the surface was then dried with air flow. This sample preparation method was used to eliminate any remaining salts and minimize artifact aggregation during drying. The measurements were made on various substrates. We used mica, highly oriented pyrolytic graphite (HOPG), and hexamethyldisilazane-modified mica (HMDS-mica). The surface of freshly cleaved mica was rendered hydrophobic using HMDS as the surface modification agent. Pieces of mica were kept in an HMDS atmosphere for 10 h in a vacuum desiccator and were then ready for use. For experiments on clot formation, a small amount of thrombin was added to the fibrinogen solution. In the major part of the experiments, we used 0.2 μL of thrombin (final concentration 3.6 nM, specific activity 1.4 kIU/mg) and 20 μL of fibrinogen solution of different concentrations. To obtain a large network of fibers, fibers with molecular resolution, or thin single-stranded associates and fibrinogen molecules, 9 μM, 200 nM, or 20 nM fibrinogen solutions were, respectively, used. Strong interaction was noticed between fibrinogen and HOPG in control experiments (Figure 1D). To avoid this additional interaction all samples for experiments on clot formation were made on only two surfaces: mica and HMDS-mica. 4923
dx.doi.org/10.1021/la200148n |Langmuir 2011, 27, 4922–4927
Langmuir
ARTICLE
Figure 2. Height mode AFM images of the fibrin network on mica: 30 s after thrombin was added to a 9 μM fibrinogen solution, Z range 45 nm (A); 510 min after thrombin was added to a 9 μM fibrinogen solution, Z range 140 nm (B); and 1 min after thrombin was added to a 0.2 μM fibrinogen solution Z range 55 nm (C) and Z range 12 nm (D). After thrombin is added to the fibrinogen solution, the clotting process proceeds very quickly and irreversibly. Preparing samples with high and medium densities therefore requires shorter exposure times. Thrombin was usually added and immediately applied a drop on the substrate surface for 1, 2, or 5 min. To prepare the samples with low fibrinogen concentrations, a mixture was kept in solution for 5 min and then on the substrate surface for another 5 min. Each experiment was conducted at least 5 times. Different cantilevers were used to minimize AFM tip artifacts. The entire set of data was used to calculate the means values and standard deviations of heights, lengths, and other parameters. Image Processing. The FemtoScan Online software (ATC, Russia) was used to analyze and present AFM data. Images were typically flattened with a linear or parabolic fit and averaged by lines. Algorithms of the actual surface profile recovering were not used. The widths of objects measured in the XY plane have a certain broadening.
’ RESULTS AND DISCUSSION Single Fibrinogen Molecules in AFM. A series of control experiments with fibrinogen was conducted to determine the most suitable substrate for investigating the clotting process further. Mica was previously used as a substrate for fibrinogen in AFM studies,10,11,13,14,16 and our data confirmed that fibrinogen molecules retained their domain structure on it. Although fibrinogen had a clear three-domain structure in electron microscopy experiments,5,6,19 in AFM, fibrinogen molecules can be visualized in compact form as globules11,13 or in extended form with a visible domain structure.10,11,14,16 Such a different appearance of the molecules is primarily associated with the fact that different
cantilevers were used. In addition, fibrinogen is a very flexible molecule that can take a variety of 2D conformations when adsorbed to the surface. In our case the domain structure was visible on mica but only 20% of the fibrinogen molecules had an apparent three-domain structure and about 80% of the molecules looked like two domains (dumbbells). Both types of molecules are shown in Figure 1A and 1B. Possible location of fibrinogen domains on the substrate surface can be supposed from zoomed images (Figure 1B). It is clearly seen that the E domain is distinguishable in elongated molecules only. Therefore, the dumbbell-like fibrinogen structure in AFM experiments is a result of an apparent broadening due to a cantilever tip radius that is larger than the protein diameter. The average height of the D domain on mica is 2.2 ( 0.4 nm, and the distance between the two end domains is 20 ( 4 nm. The height of the central E domain is 1.1 ( 0.5 nm, and the average length of the molecule is 42 ( 7 nm. Images obtained on HMDS-mica are completely different. Because this surface is hydrophobic, the molecules spread on the surface and almost lose their domain structure (Figure 1C). The average height of the observed objects is 1.2 ( 0.4 nm. On HOPG, the images again differ. The HOPG surface is highly hydrophobic and has many steps. As shown in Figure 1D, fibrinogen molecules on HOPG are stretched and tend to aggregate into large structures oriented in the step direction. The average height of these aggregates is 2.0 ( 0.5 nm. This result agrees well with previously obtained data.13,14 Mica was concluded to be the best substrate because fibrinogen molecules retain their domain structure. 4924
dx.doi.org/10.1021/la200148n |Langmuir 2011, 27, 4922–4927
Langmuir
ARTICLE
Figure 3. Height mode AFM images of early fibrils of fibrin on mica, Z range 6 nm (A, C, and E), and on HMDS-mica, Z range 5 nm (B, D, and F).
AFM Investigation of Thrombin-Induced Fibrin Fiber Formation. It is extremely inconvenient to study fiber network
formation using such a fibrinogen concentration as in human blood. Solution of 9 μM fibrinogen was used, and a network of large fibers was formed on mica in 30 s after thrombin was added (Figure 2A). The height of these structures is 11.7 ( 4.5 nm. In 510 min the surface is covered with the web (Figure 2B), which we suppose is a rigid clot. Any details are hardly visible under these conditions; we can characterize the surface of the clot only by its roughness. The root-mean-square roughness is about 30 nm. Decreasing the fibrinogen concentration down to 0.2 μM we obtained better resolution and could see the molecular period on fibers 1 min after thrombin was added. These images are shown in Figure 2C and 2D. Under these conditions, the fibers are also long but slightly thinner: the height is 8.5 ( 2.5 nm. The most interesting feature is a 23 nm period, which nicely agrees with classical electron microscopy results.4
AFM Investigation of Early Fibrin Associates. The next series of AFM experiments was made with the fibrinogen concentration further decreased to 20 nM. Under these conditions, molecularly thin fibrin associates about 1 μm long and one or a few fibrin molecules wide were observed. Surprisingly, the structure of the main part of the associates differed from the protofibril structure. We observed the same behavior on both mica and HMDS-mica. Typical images obtained on pure mica are shown in Figure 3A, 3C, and 3E. Two types of objects can be seen in these images: double-stranded (protofibril) and single-stranded (coined as ‘profibril’) associates. There are also large associates that have the height of a single molecule and are composed of profibrils and protofibrils (Figure 3A and 3C). The image resolution allowed measuring the geometric parameters of both types of associates. The average protofibril height is 3.0 ( 0.3 nm, which is slightly higher than the height of the D domain in fibrinogen molecules. 4925
dx.doi.org/10.1021/la200148n |Langmuir 2011, 27, 4922–4927
Langmuir
ARTICLE
Figure 4. Volume distribution of fibrin associates from a 20 nM fibrinogen solution at 0 (red), 1 (green), 5 (blue), and 10 min (magenta) after thrombin addition.
Table 1. Hydrodynamic Radii from DLS Data (size distribution by volume is shown in parentheses)
low-weight particles high-weight particles
without thrombin
1 min reaction
5 min reaction
10 min reaction
13.2 nm (100%)
13.2 nm (93%) 87 nm (7%)
23 nm (58%) 350 nm (42%)
27 nm (54%) 380 nm (46%)
Figure 5. Possible mechanisms of fibrin associates formation (fibrin E domains are shown in brown; D domains are shown in yellow): conventional protofibril formation via DED interactions (A), fibrin molecules are fastened together through two D domains directly (B), through RC-domain dimerization (C), or through both DD and RC domains interactions (D).
The protofibril width in AFM is about 27 nm. This seems reasonable because the real diameter of the protofibril structure should be about 12 nm, and the typical tip radius of the cantilever is between 6 and 10 nm. Taking a tip radius equal to 8 nm we obtain a width of 12 þ 7 2 = 26 nm for these associates in the AFM experiment (Figure 3E). The broadening was derived from a simple geometric model similar to the model used in ref 19. On the other hand, the width of profibrils, which are the most interesting structures, is much less, 19 nm. This is close to the estimated value of the fibrinogen diameter in AFM 6 þ 2 7 = 20 nm; therefore, profibrils are single stranded. The average profibril height is 2.8 ( 0.3 nm, which is close to the D domain height. There are also gaps between profibril domains, and these gaps could be spaces between the D domains of either different fibrin molecules or the same molecule (or both cases simultaneously). Unfortunately, AFM cannot distinguish between these two hypotheses. The average distance between the centers of two neighboring domains in profibrils is 22 ( 9 nm. The significant deviation from the mean may occur because the distribution in fact has two peaks (although not very well distinguishable) or because the molecules in profibrils are connected with a flexible bond.
We conducted the same experiments on HMDS-mica (Figure 3B, 3D, and 3F) to exclude substrate-induced profibril formation. On this surface, fibrinogen is spread-eagled and the overall structures looked slightly lower and thinner (on HMDS-mica, the average protofibril height is 2.8 ( 0.3 nm and the width is 20.3 ( 3.5 nm, while the average profibril height is 2.7 ( 0.3 nm and the width is 14.3 ( 2.4 nm). An interesting detail becomes evident from comparing Figures 1C and 3F: on HMDS-mica, fibrinogen molecules lost their domain structure but profibrils retained it and have extremely smooth gaps. This indicates that the profibril structure is more stable than the fibrinogen structure on a hydrophobic substrate and could hardly be assembled from fibrin molecules on the substrate surface. This indicates that the profibril structure is more stable than the fibrinogen structure on a hydrophobic substrate, and protofibrils could hardly either be assembled from fibrin molecules on the substrate surface or be an artifact of sample drying. DLS Experiments. Fibrin association from 20 nM fibrinogen solution had not been previously demonstrated. We therefore performed DLS experiments to confirm the correspondence between the bulk-solution and near-surface processes. We also analyzed the 2 and 0.2 μM fibrinogen solutions. The estimated 4926
dx.doi.org/10.1021/la200148n |Langmuir 2011, 27, 4922–4927
Langmuir hydrodynamic radius for a fibrinogen sample is 13.2 ( 0.3 nm (Figure 4), which agrees well with the hydrodynamic radius of 12.7 ( 0.6 nm determined by Wasilewska et al.20 Thrombin-induced association was observed for all fibrinogen concentrations. The largest fibrils were formed from the 2 μM fibrinogen sample (the hydrodynamic radius was about 18 μM after 5 min). A considerable fraction (2030% by volume) of fibrinogen, fibrin, and early oligomers was observed in the 0.2 μM fibrinogen samples up to 5 min after thrombin was added, and the hydrodynamic radius was about 1.9 μm, which agrees well with Figure 2C. The least association degree was in the 20 nM fibrinogen samples: 50% at 510 min after the reaction start (Table 1 and Figure 4). This trend is also reflected in the AFM data (Figure 3). We can conclude that AFM images rather reliably reflect fibrin association in a bulk solution.
’ CONCLUSIONS The series of AFM experiments were performed to investigate different stages of fibrin association in vitro. Using a physiological fibrinogen concentration results in web formation, and any details with molecular resolution are indistinguishable in the web. Decreasing of the fibrinogen concentration down to 200 nM led to the appearance of a distinct pattern on the fiber surface: the 23 nm period was observed, which agrees with classical electron microscopy results. Decreasing the fibrinogen concentration further down to 20 nM led to highly resolved images of early fibrin associates: classical form, protofibrils, and a new form, profibrils. The conventional fiber association mechanism assumes formation of a double-stranded form, protofibril, through DED interactions (Figure 5A). The single-stranded associates discovered here, profibrils, clearly have a domain structure, and the geometric parameters of those domains are equal to the fibrinogen domain parameters. Using two different substrates (mica and HMDS-mica) with different hydrophilic properties was shown to have no effect on profibril formation. Although a considerable part of fibrinogen molecules lost their domain structure on HMDS-mica, profibrils retained their multidomain pattern, and these domains looked much more compact than those of fibrinogen on HMDS-mica. This effect did not change with sorption duration. Finally, the association in solution was directly shown by the DLS experiments. Taken altogether these results indicate profibril formation in a bulk solution. Some possibilities for profibril formation are shown schematically in Figure 5. On the basis of the background described in the Introduction, the following mechanisms of profibril formation can be proposed: DD interactions (Figure 5B), RC domain dimerization (Figure 5C), and a hybrid of DD and RC interactions (Figure 5D). The detailed mechanism of initial association must nevertheless be studied more thoroughly.
ARTICLE
(2) Yakovlev, S.; Mosesson, M. W.; Gilliland, G. L.; Medved, L.; Pechik, I. Biochemistry 2006, 45, 3588–3597. (3) Yang, Z.; Mochalkin, I.; Doolittle, R. F. Proc. Natl. Acad. Sci. U.S. A. 2000, 97, 14156–14161. (4) Williams, R. C. Proc. Natl. Acad. Sci. U.S.A. 1983, 80, 1570–1573. (5) Fowler, W. E.; Hantgan, R. R.; Hermans, J.; Erickson, H. P. Proc. Natl. Acad. Sci. U.S.A. 1981, 78, 4872–4876. (6) Veklich, Yu. I.; Gorkun, O. V.; Medved, L. V.; Nieuwenhuizen, W.; Weisel, J. W. J. Biol. Chem. 1993, 268, 13577–13585. (7) Mosesson, M. W.; DiOrio, J. P.; Siebenlist, K. R.; Wall, J. S.; Hainfeld, J. F. Blood 1993, 82, 1517–1552. (8) Brown, A. E. X.; Litvinov, R. I.; Discher, D. E.; Wiesel, J. W. Biophys. J. 2007, L, 39–41. (9) Koo, J.; Rafailovich, M. H.; Medved, L.; Tsurupa, G.; Kudryk, B. J.; Liu, Y.; Galanakis, D. K. J. Thromb. Haemost. 2010, 8, 2727–35. (10) Abou-Saleh, R. H.; Connell, S. D.; Harrand, R.; Ajjan, R. A.; Mosesson, M. W.; Smith, D. A. M.; Grant, P. J.; Ariens, R. A. S. Biophys. J. 2009, 96, 2415–2427. (11) Tsapikouni, T. S.; Missirlis, Y. F. Colloids Surf., B: Biointerfaces 2007, 57, 89–96. (12) Cacciafesta, P.; Humphris, A. D. L.; Jandt, K. D.; Miles, M. J. Langmuir 2000, 16, 8167–8175. (13) Marchin, K. L.; Berrie, C. L. Langmuir 2003, 19, 9883–9888. (14) Agnihotri, A.; Siedlecki, C. A. Langmuir 2009, 20, 8846–8852. (15) Yaseen, M.; Zhao, X.; Freund, A.; Seifalian, A. M.; Lu, J. R. Biomaterials 2010, 31, 3781–3792. (16) Adamczyk, Z.; Nattich, M.; Wasilewska, M.; Sadowska, M. J. Colloid Interface Sci. 2011in press. (17) O’Brien, E. T.; Falvo, M. R.; Millard, D.; Eastwood, B.; Taylor, R. M.; Superfine, R. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 19438–19443. (18) Litvinov, R. I.; Yakovlev, S.; Tsurupa, G.; Gorkun, O. V.; Medved, L.; Weisel, J. W. Biochemistry 2007, 46, 9133–9142. (19) Santos, N. C.; Ter-Ovanesyan, E.; Zasadzinski, J. A.; Prieto, M.; Castanho, M. A. R. B. Biophys. J. 1998, 75, 1869–1873. (20) Wasilewska, M.; Adamczyk, Z.; Jachimska, B. Langmuir 2009, 25, 3698–3704.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected],
[email protected].
’ REFERENCES (1) Pechik, I.; Madrazo, J.; Mosesson, M. W.; Hernandez, I.; Gilliland, G. L.; Medved, L. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 2718–2723. 4927
dx.doi.org/10.1021/la200148n |Langmuir 2011, 27, 4922–4927
