http://informahealthcare.com/plt ISSN: 0953-7104 (print), 1369-1635 (electronic) Platelets, Early Online: 1–6 ! 2014 Informa UK Ltd. DOI: 10.3109/09537104.2014.940888
ORIGINAL ARTICLE
Use of non-contact hopping probe ion conductance microscopy to investigate dynamic morphology of live platelets Xiao Liu1,2, Ying Li1, Hui Zhu1, Zilong Zhao1, Yuan Zhou1, Ana-Maria Zaske3, Li Liu1, Min Li4, Hujie Lu1,2, Wei Liu1, Jing-Fei Dong5,6, Jianning Zhang1, & Yanjun Zhang1,2,7
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1
Key Laboratory of Post-trauma Neuro-repair and Regeneration in Central Nervous System, Ministry of Education, Tianjin Key Laboratory of Injuries, Variations and Regeneration of Nervous System, Tianjin Medical University General Hospital, Tianjin Neurological Institute, Tianjin, China, 2 Nanomedicine Laboratory, China National Academy of Nanotechnology & Engineering, Tianjin, China, 3Division of Cardiology, Department of Internal Medicine, the University of Texas Houston Health Science Center at Houston, Houston, TX, USA, 4Pathology Institute, School of Basic Medical Sciences, Lanzhou University, Lanzhou, China, 5Puget Sound Blood Center, Seattle, WA, USA, 6Division of Hematology, Department of Medicine, University of Washington, School of Medicine, Seattle, WA, USA, and 7Division of Medicine, Imperial College London, London, UK Abstract
Keywords
Circulating platelets are anucleated and multi-functional cells that participate in hemostasis and arterial thrombosis. Multiple ligands and mechanical forces activate platelets, leading to cytoskeletal rearrangement and dramatic shape-changes. Such dramatic changes in platelets membrane structures are commonly detected by optical and electron microscopy after platelets are fixed. We have recently developed a method to study the membrane morphology of live platelets using Hopping Probe Ion Conductance Microscopy (HPICM). We have successfully used this technology to study the process of platelet microvesiculation upon exposure to selective agonists. Here, we further discussed technical details of using HPICM to study platelet biology and compared results from HPICM to those from conventional atomic force microscopy and scanning electron microscopy. This method offers several advantages over current technologies. First, it monitors morphological changes of platelets in response to agonists in real time. Second, platelets can be repeatedly scanned over time without damages brought by heat and prolong light exposure. Third, there is no direct contact with platelet surface so that there will no or minimal mechanical damages brought by a cantilever of a conventional atomic force microscopy. Finally, it offers the potential to study platelet membrane ion channels, which have been technically challenging up-to-date. Our data show that HPICM has high-resolution in delineating changes of platelet morphology in response to stimulations and could help to unravel the complex role of platelet in thrombus formation.
AFM, HPICM, SICM, SPM
Introduction Circulating platelets are anucleated and multifunctional cells that are active in hemostasis and arterial thrombosis [1]. Multiple ligands and mechanical forces activate platelets, leading to cytoskeletal rearrangement and dramatic changes in platelet morphologies [2–4]. However, even with significantly improved resolution, a conventional light microscope is insufficient to study these morphological changes and their biological significance. A high-resolution scanning electron microscopy (SEM) is used to study platelet surface and intracellular structures [5], but it requires platelets to be fixed, and, therefore, is very limited in studying dynamical changes of platelet morphology induced by a ligand. Dehydration, fixation, and stained with heavy metal particles may also alter the morphology of a platelet [5, 6]. Confocal microscopy has been increasingly used to monitor
Correspondence: Yanjun Zhang, Tianjin Neurological Institute, Tianjin Medical University General Hospital, 154 Anshan Road, Tianjin 300052, China. Tel: +86 22 60817460. E-mail: [email protected] Jianning Zhang, Department of Neurosurgery, Tianjin Medical University General Hospital, 154 Anshan Road, Tianjin 300052, China. Tel: +86 22 60362026. E-mail: [email protected]
History Received 23 May 2014 Revised 27 June 2014 Accepted 30 June 2014 Published online 6 August 2014
platelet response to specific ligands by allowing repeat scanning platelets over a long period of time without fixation [7]. However, this technique is limited in resolution to delineate fine 3D membrane structures of platelet and subjected platelets to photo-bleaching and photo-toxicity due to a prolonged light exposure [8]. Compared to a conventional microscopy that ‘‘sees’’ a platelet, scanning probe microscopy (SPM) is increasingly being used to study the structure and functions of a cell by ‘‘touching’’ [9]. Atomic force microscopy (AFM) is a representative of SPM and defines non-conducting surface by ‘‘touching’’ cells in culture [10]. However, platelets are highly sensitive to mechanical forces generated by an AFM probe [7, 10], which could produce a lateral shear force between the cantilever tip and platelet surface. This force could be sufficient to disrupt platelet’s membrane or alter its functions during repetitive scans [11, 12]. Scanning ion conductance microscopy (SICM) overcomes some of these technical obstacles [13–18], while produces high-resolution topographic images of platelets without fixation and directly contact [19–21]. However, the inherent delay in a feedback response of SICM can be disruptive [22], making it difficult to follow abrupt upward/ downward steps during dynamic morphological changes of live cells. Our recent observation suggests that hopping probe ion
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conductance microscopy (HPICM) overcomes these limitations to produce high temporal resolutions of platelets without directly contacting them [22–25]. Here, we discuss technical aspects and potential applications of HPICM in studying live platelets.
Materials and methods
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Preparation of platelets for imaging Human blood samples were obtained from healthy subjects using sodium citrate as anticoagulant (0.38% final concentration) under a protocol approved by the ethics committee of Tianjin Medical University General Hospital. Platelet-rich plasma (PRP) was obtained by centrifuging whole blood at 150 g for 15 minutes at room temperature and used in all experiments. Polystyrene cell culture dishes (Corning, USA) were coated with either human fibrinogen (1 mg/mL, Hematologic Technologies, Essex Junction, VT) or poly-D-lysine (10 mg/mL, MW 70–150 kDa, Sigma-Aldrich, St. Louis, MO) overnight at 4 C. Platelets were then allowed to adhere to fibrinogen or polyD-lysine for 30 minutes at 37 C. Fibrinogen binds the platelet integrin aIIbb3, whereas poly-D-lysine promotes platelet adhesion through electrostatic interactions. In the end of the incubation, platelets were washed thrice with PBS and twice times with L15 medium (Gibco, Langley, OK) to remove unbound platelets. Platelet imaging by hopping probe ion conductance microscopy Hopping Probe Ion Conductance Microscopy (HPICM) system consisted of an ICnano scanner controller (Ionscope Ltd, UK) and a scan head SH01 (Ionscope Ltd, UK), as described previously [22, 24]. The SH01 scan head with a nanopipette was placed on the platform of an inverted TiU microscope (Nikon Corporation, Tokyo, Japan). The nanopipette was made by pulling borosilicate glass (O.D. 1.00 mm, I.D. 0.59 mm, VitalSense Scientific Instruments Co., China) on a P-2000 laser-based puller (Sutter Instruments Co., Novato, CA) in two pulling cycles: (1) HEAT 340, FIL 4, VEL 40, DEL 200 and PUL 0, and (2) HEAT 340, FIL 3, VEL 30, DEL 180 and PUL 250. The resistance of an HPICM nanopipette was approximately 130 M when it was filled with L15 medium. Inner and the outer diameters of the nanopipettes were 75 nm and 112 nm, respectively, as evaluated by SEM [25]. This nanopipette had an inner radius of about 30 nm inner radius and allowed a maximum spatial resolution of 30 nm [26]. An external Axon MultiClamp 700B amplifier (Molecular Devices, Sunnyvale, CA) was connected to the nanopipette electrode to supply a DC voltage of +200 mV, and the ionic current flowing into the nanopipette was 1.5 nA. HPICM controlled a vertical Z direction Piezo to allow this nanopipette to hop over the surface of platelets. A 0.3–0.4% reduction of the ion-current flowing into nanopipette was set to maintain a constant separation between the nanopipette and platelet surface. During scan, the nanopipette performed pre-scan hopping distance at 3000 nm and 400 nm amplitude for high-resolution scanning. The nanopipette falling rate was set to 50 nm/ms, meanwhile its rising rate was increased to 500 nm/ms. An adaptive imaging protocol of HPICM was used for scanning platelets [22]. Briefly, HPICM pre-scanned a designated area to estimate the overall roughness by measuring the difference in heights at the four corners. If a platelet was detected in the area, HPICM subsequently scanned this platelet at a higher resolution of 256 256 pixels. In contrast, HPICM scanned a flat surface identified in the pre-scan at a lower resolution of 128 128 pixels. The operation time of this adaptive scanning was further reduced by obtaining high-resolution scan immediately after the pre-scan at the four corner points. This is particularly useful for
Platelets, Early Online: 1–6
imaging living platelets because it ensures that platelets undergo minimal shape changes between pre-scan and high-resolution scan. This adaptive operation protocol also accelerated the speed of HPICM scan without the loss of high-resolutions. At this setting, each scan took approximately 6–9 min. During the entire scan period, platelets were bathed in L15 medium and maintained at 24–28 C. Acquired raw topography data were bilinearinterpolated using ScanIC Image Viewer version 1.0 (Ionscope Ltd, Royston, UK) to produce final images of 512 512 pixels. Platelet imaging by AFM For comparison, human platelets were scanned by AFM. AFM was performed in liquid at a rate of 0.3 Hz in a contact mode with DNP-S cantilevers (fo ¼ 12–24 kHz, k ¼ 0.06 N/m, Bruker Corporation, Santa Barbara, CA) using a BioScope IITM Controller (Bruker Corporation) integrated into a Nikon TE2000-E inverted optical microscope (Nikon Instruments, Inc. Lewisville, TX) as previously described [27]. The graphic program Research NanoScope version 7.3 was used to convert digital data into topographical images. The time required to scan a 50 50 mm2 area with a typical resolution of 512 512 pixels by AFM was approximately 26 min. Platelet imaging by SEM For comparison, human platelets were also scanned with SEM. Carbon grids were coated with fibrinogen over night at 4 C and then incubated with PRP for 30 minutes at 37 C. After nonadherent platelets were removed by washing, platelets on grids were fixed with 3% of glutaraldehyde, dehydrated with increasing concentrations of ethanol (70%, 90% and 100%), and rehydrated with tert-butyl alcohol as previously discussed [27]. SEM micrographs were recorded with JSM-6380Lv SEM (JEOL Ltd., Tokyo, Japan). Statistical analysis HPICM images were processed and analyzed by ScanIC Image Viewer version 1.0 program (Ionscope Ltd, UK). All data were analyzed using Origin 8.0 program (Originlab Corporation, Northampton, MA). Quantitative values were presented as mean ± SEM. All data were analyzed by pair comparison. A difference between means at the level of p50.05 was considered statistically significant; p50.005 was statistical highly significant.
Results Continuous HPICM observations of live platelets With technical settings discussed in the ‘‘Method’’ section, we were able to detect live platelets adherent to fibrinogen at a spatial resolution of 30 nm, without direct contacts between platelets and the nanopipette. Platelets remained adherent and stable over a 60–90 minutes period of repeated scans (6–10 cycles) by HPICM [27]. Figure 1 shows HPICM topographical images of platelets adherent to fibrinogen taken every 9 minutes. Without exposure to an agonist, platelets adherent to fibrinogen presented in two distinct morphologies: low-density spread shape (LDSS) and high-density bubble shape platelets (HDBS) (note morphological changes of six platelets highlighted by white dotted ellipses). This difference in density indicates the different heights of adherent LDSS (0.51 ± 0.13 mm, n ¼ 35) and HDBS (1.88 ± 0.54 mm, n ¼ 36) platelets. In addition, LDSS platelets were larger (5.68 ± 1.44 mm, n ¼ 56) than HDBS platelets (2.95 ± 0.62 mm, n ¼ 72), suggesting that LDSS platelets spread completely on fibrinogen. Some HDBS platelets gradually
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DOI: 10.3109/09537104.2014.940888
Morphology imaging of live platelet by HPICM
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Figure 1. Repetitive HPICM scans of live platelets on fibrinogen substrate. Six images were acquired every 9 minutes over a 45 minutes time-lapse scanning of a selected 50 50 mm area.
Figure 2. High-resolution HPICM images of changed morphology from HDBS to LDSS. (A) Three images constituted a sequence of 8 minutes time-lapse scanning. (B) Scanning profile changes of the solid line via pseudo-nuclear center in A from 0 minutes (thick black line trace) to 8 minutes (grey line trace) to 16 minutes (thin black line trace).
became a LDSS morphology (Figure 1, arrow), while others remained unchanged, but with slightly shifted their positions (white solid ellipse) during repeated scans. In contrast, LDSS platelets underwent little change in morphology during the same period. These data suggest that the membranes of live platelets are highly dynamic and the two distinct morphologies of LDSS and HDBS likely represent two interchangeable stages of platelet adhesion to and spreading on fibrinogen. One set of high-resolution time-lapsed HPICM scans detected this gradual change in morphology from HDBS to LDSS in approximately 16 minutes (Figure 2). During spreading, the height of HDBS platelets at the pseudo-nuclear center reduced from 1.74 ± 0.50 mm to 0.51 ± 0.18 and its diameter increased from 2.66 ± 0.41 mm to 5.33 ± 1.10 mm, leading to an increase in the surface area from 14.66 ± 4.99 mm2 to 28.70 ± 10.95 mm2 without significant change in volume (n ¼ 11). A scanning profile of the platelet cross-section marked by the straight-line across through three time-lapse frames show such morphological changes during the scanning period (Figure 2B). These data for the first time showed platelet membrane fluidity and spreading in high resolution. Because of its ability to repeatedly scan live platelets with no direct light and heat exposure, HPICM is ideally to observe the dynamic process of platelet spreading on different matrixes. Platelets were allowed to adhere to either human fibrinogen, which is mediated by the platelet integrin aIIbb3 [28,29], or
positively charged poly-D-lysine, which mediated platelet adhesion through an electrostatic interaction [30]. Both HDBS and LDSS morphologies were also observed from those adherent platelets on poly-D-lysine (Figure 3E–H). However, platelets on poly-D-lysine were significantly larger and thicker (with a big volume) than those on fibrinogen. Both pseudo-nuclear center height and cell volume of HDBS platelets on poly-D-lysine were significantly larger than those on human fibrinogen (Table I). Morphology of platelets observed by SEM and AFM We compared morphological characteristics of adherent platelets on fibrinogen observed by HPICM to those detected by AFM and SEM (Figure 4). An AFM cantilever probes platelets by touching their surfaces and was able to detected HDBS and LDSS morphologies of adherent platelets [27]. However, an AFM cantilever removed a substantial number of adherent platelets (35%) during repeat scans (data not shown), generating different ratios of HDBS to LDSS platelets on fibrinogen (0.21 ± 0.03 for AFM, n ¼ 6, vs. 0.94 ± 0.54 for HPICM, n ¼ 10). This observation suggests that the mechanical force loading on platelets by the cantilever may have pulled platelets from the substrate and HDBS platelets were more sensitive to this mechanical pulling force. To address this concern, we also scanned platelets fixed by paraformaldehyde (Figure 4B). The fixation significantly reduced the number of platelets that were removed by the cantilever, but
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Platelets, Early Online: 1–6
Figure 3. HPICM images of HDBS and LDSS morphologies of live human platelets adherent to fibrinogen (A–D) and poly-D-lysine (E–H). B–D and F–H are gradually zoom-in scans of HDBS and LDSS platelets corresponding to those white-dotted-squares marked areas of B–D and F–H in figures A–B and E–F, respectively. Table I. Measurements of platelets adherent to fibrinogen and poly-D-lysine.
HDBS (n ¼ 36) Height (mm) Volume (mm3) LDSS (n ¼ 35) Height at peudo-center (mm) Volume (mm3)
Fibrinogen
Poly-D-lysin
p Value
1.88 ± 0.54 11.00 ± 3.75 0.51 ± 0.13 13.44 ± 4.47
2.63 ± 0.65 17.02 ± 5.90 0.89 ± 0.45 18.23 ± 11.57
50.005 50.005 50.005 50.05
Figure 4. Platelets adherent to fibrinogen-coated surface imaged by SEM (A) and AFM (B). Arrows pointed platelets had low density spread shape (LDSS) and circle surrounded platelets had high density bubble shape (HDBS).
some of pseudo-center of HDBS platelets were scratched by the cantilever because the distance between the cantilever and platelet is fixed. Platelets adherent to fibrinogen were also fixed and imaged by SEM (Figure 4A). Both HDBS and LDSS platelets were identified at a ratio similar to that of HPICM. However, both AFM and SEM were unsuitable to detect dynamic changes in living platelet morphologies.
Discussion We have shown that HPICM is an effective tool in monitoring morphological changes of adherent platelets in real-time. Using this HPICM technology, we identified two distinct morphologies of live platelets adherent to fibrinogen: fully spread (LDSS) and
non-spreading (HDBS). These morphologies are highly dynamic in real-time and HDBS platelets gradually became LDSS morphology in 10–20 minutes. These two morphologies were also observed from platelets adhered to poly-D-lysine, suggesting that their formation is independent of the interaction between fibrinogen and the platelet integrin aIIbb3. However, this morphological distinct is important because we have recently shown that HDBS, but not LDSS, platelets are very sensitive to agonist-induced microvesiculation [27]. These dynamic and heterogeneous characteristics of platelets are distinct from neurons and epithelial cells that have previously been studied with this technology [22–25]. A key technical feature of HPICM is its hopping probe, which provides a sensitive tip-sample separation control to perform non-contact surface profiling [21–25] and will not disturb the
Morphology imaging of live platelet by HPICM
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DOI: 10.3109/09537104.2014.940888
platelet membrane. In contrast, a conventional AFM requires a suitable surface viscosity and stiffness of a cell. Because the distance between a surface and a cantilever is fixed, AFM is less flexible in probing a cell that undergoes morphological changes. For example, a setting for tall HDBS platelets may sacrifice the resolution for LDSS platelets, whereas the tip of AFM cantilever could scratch the membrane surface at the top of a HDBS platelet when the distance was set for fully spread platelets, resulting in image distortion as seen in Figure 4(B) (highlighted by white dotted circle). Another drawback for AFM imaging is that an acquisition time needs to be increased when a dynamic process is to be visualized. High-speed AFM has been recently used to directly visualize the dynamics structure and processes of biological molecules in physiological solutions [31–33], but its application in live cells is few. This technical obstacle of a longer acquisition time is also addressed by HPICM, which uses an adaptive resolution scanning protocol to reduce the acquisition time for one frame of scanning of living platelets in a 50 50 mm2 area to 6–9 minutes. In conclusion, upon binding to the subendothelial matrix, platelets are activated and undergo drastic shape changes that are critical for platelet adhesion and aggregation [34–36]. These morphological changes of platelets have been widely reported, but are mostly observed after platelets were fixed with a chemical fixative. HPICM overcomes several key technical difficulties associated with confocal microscopy, AFM and SEM to offer high-resolution monitoring morphological changes and defining their physiological significance in real-time without inducing mechanical and chemical damages to platelets. Time lapse HPICM scans offer a unique opportunity to monitor the agonist-induced dynamic changes in platelet morphologies that have been difficult to detect with other imaging technologies [27]. HPICM can also be combined with patch-clamping technology using the same nanopipette [24, 25], so that one can simultaneously monitor platelet shape changes and the activity of ion channels during platelet activation, which has not been possible with current imaging technologies.
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Declaration of interest This work was supported by Tianjin Science and Technology Support Program of China (No.14ZCZDSY00020), Tianjin Natural Science Foundation of China (No.13JCYBJC21900 and 12JCYBJC31500), National Natural Science Foundation of China (No.31300828, 81330029, and 81271361), and the National Heart, Lung, and Blood Institute, NIH, USA (grants HL71895 and HL85769).
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ORIGINAL ARTICLE
Use of non-contact hopping probe ion conductance microscopy to investigate dynamic morphology of live platelets Xiao Liu1,2, Ying Li1, Hui Zhu1, Zilong Zhao1, Yuan Zhou1, Ana-Maria Zaske3, Li Liu1, Min Li4, Hujie Lu1,2, Wei Liu1, Jing-Fei Dong5,6, Jianning Zhang1, & Yanjun Zhang1,2,7
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1
Key Laboratory of Post-trauma Neuro-repair and Regeneration in Central Nervous System, Ministry of Education, Tianjin Key Laboratory of Injuries, Variations and Regeneration of Nervous System, Tianjin Medical University General Hospital, Tianjin Neurological Institute, Tianjin, China, 2 Nanomedicine Laboratory, China National Academy of Nanotechnology & Engineering, Tianjin, China, 3Division of Cardiology, Department of Internal Medicine, the University of Texas Houston Health Science Center at Houston, Houston, TX, USA, 4Pathology Institute, School of Basic Medical Sciences, Lanzhou University, Lanzhou, China, 5Puget Sound Blood Center, Seattle, WA, USA, 6Division of Hematology, Department of Medicine, University of Washington, School of Medicine, Seattle, WA, USA, and 7Division of Medicine, Imperial College London, London, UK Abstract
Keywords
Circulating platelets are anucleated and multi-functional cells that participate in hemostasis and arterial thrombosis. Multiple ligands and mechanical forces activate platelets, leading to cytoskeletal rearrangement and dramatic shape-changes. Such dramatic changes in platelets membrane structures are commonly detected by optical and electron microscopy after platelets are fixed. We have recently developed a method to study the membrane morphology of live platelets using Hopping Probe Ion Conductance Microscopy (HPICM). We have successfully used this technology to study the process of platelet microvesiculation upon exposure to selective agonists. Here, we further discussed technical details of using HPICM to study platelet biology and compared results from HPICM to those from conventional atomic force microscopy and scanning electron microscopy. This method offers several advantages over current technologies. First, it monitors morphological changes of platelets in response to agonists in real time. Second, platelets can be repeatedly scanned over time without damages brought by heat and prolong light exposure. Third, there is no direct contact with platelet surface so that there will no or minimal mechanical damages brought by a cantilever of a conventional atomic force microscopy. Finally, it offers the potential to study platelet membrane ion channels, which have been technically challenging up-to-date. Our data show that HPICM has high-resolution in delineating changes of platelet morphology in response to stimulations and could help to unravel the complex role of platelet in thrombus formation.
AFM, HPICM, SICM, SPM
Introduction Circulating platelets are anucleated and multifunctional cells that are active in hemostasis and arterial thrombosis [1]. Multiple ligands and mechanical forces activate platelets, leading to cytoskeletal rearrangement and dramatic changes in platelet morphologies [2–4]. However, even with significantly improved resolution, a conventional light microscope is insufficient to study these morphological changes and their biological significance. A high-resolution scanning electron microscopy (SEM) is used to study platelet surface and intracellular structures [5], but it requires platelets to be fixed, and, therefore, is very limited in studying dynamical changes of platelet morphology induced by a ligand. Dehydration, fixation, and stained with heavy metal particles may also alter the morphology of a platelet [5, 6]. Confocal microscopy has been increasingly used to monitor
Correspondence: Yanjun Zhang, Tianjin Neurological Institute, Tianjin Medical University General Hospital, 154 Anshan Road, Tianjin 300052, China. Tel: +86 22 60817460. E-mail: [email protected] Jianning Zhang, Department of Neurosurgery, Tianjin Medical University General Hospital, 154 Anshan Road, Tianjin 300052, China. Tel: +86 22 60362026. E-mail: [email protected]
History Received 23 May 2014 Revised 27 June 2014 Accepted 30 June 2014 Published online 6 August 2014
platelet response to specific ligands by allowing repeat scanning platelets over a long period of time without fixation [7]. However, this technique is limited in resolution to delineate fine 3D membrane structures of platelet and subjected platelets to photo-bleaching and photo-toxicity due to a prolonged light exposure [8]. Compared to a conventional microscopy that ‘‘sees’’ a platelet, scanning probe microscopy (SPM) is increasingly being used to study the structure and functions of a cell by ‘‘touching’’ [9]. Atomic force microscopy (AFM) is a representative of SPM and defines non-conducting surface by ‘‘touching’’ cells in culture [10]. However, platelets are highly sensitive to mechanical forces generated by an AFM probe [7, 10], which could produce a lateral shear force between the cantilever tip and platelet surface. This force could be sufficient to disrupt platelet’s membrane or alter its functions during repetitive scans [11, 12]. Scanning ion conductance microscopy (SICM) overcomes some of these technical obstacles [13–18], while produces high-resolution topographic images of platelets without fixation and directly contact [19–21]. However, the inherent delay in a feedback response of SICM can be disruptive [22], making it difficult to follow abrupt upward/ downward steps during dynamic morphological changes of live cells. Our recent observation suggests that hopping probe ion
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conductance microscopy (HPICM) overcomes these limitations to produce high temporal resolutions of platelets without directly contacting them [22–25]. Here, we discuss technical aspects and potential applications of HPICM in studying live platelets.
Materials and methods
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Preparation of platelets for imaging Human blood samples were obtained from healthy subjects using sodium citrate as anticoagulant (0.38% final concentration) under a protocol approved by the ethics committee of Tianjin Medical University General Hospital. Platelet-rich plasma (PRP) was obtained by centrifuging whole blood at 150 g for 15 minutes at room temperature and used in all experiments. Polystyrene cell culture dishes (Corning, USA) were coated with either human fibrinogen (1 mg/mL, Hematologic Technologies, Essex Junction, VT) or poly-D-lysine (10 mg/mL, MW 70–150 kDa, Sigma-Aldrich, St. Louis, MO) overnight at 4 C. Platelets were then allowed to adhere to fibrinogen or polyD-lysine for 30 minutes at 37 C. Fibrinogen binds the platelet integrin aIIbb3, whereas poly-D-lysine promotes platelet adhesion through electrostatic interactions. In the end of the incubation, platelets were washed thrice with PBS and twice times with L15 medium (Gibco, Langley, OK) to remove unbound platelets. Platelet imaging by hopping probe ion conductance microscopy Hopping Probe Ion Conductance Microscopy (HPICM) system consisted of an ICnano scanner controller (Ionscope Ltd, UK) and a scan head SH01 (Ionscope Ltd, UK), as described previously [22, 24]. The SH01 scan head with a nanopipette was placed on the platform of an inverted TiU microscope (Nikon Corporation, Tokyo, Japan). The nanopipette was made by pulling borosilicate glass (O.D. 1.00 mm, I.D. 0.59 mm, VitalSense Scientific Instruments Co., China) on a P-2000 laser-based puller (Sutter Instruments Co., Novato, CA) in two pulling cycles: (1) HEAT 340, FIL 4, VEL 40, DEL 200 and PUL 0, and (2) HEAT 340, FIL 3, VEL 30, DEL 180 and PUL 250. The resistance of an HPICM nanopipette was approximately 130 M when it was filled with L15 medium. Inner and the outer diameters of the nanopipettes were 75 nm and 112 nm, respectively, as evaluated by SEM [25]. This nanopipette had an inner radius of about 30 nm inner radius and allowed a maximum spatial resolution of 30 nm [26]. An external Axon MultiClamp 700B amplifier (Molecular Devices, Sunnyvale, CA) was connected to the nanopipette electrode to supply a DC voltage of +200 mV, and the ionic current flowing into the nanopipette was 1.5 nA. HPICM controlled a vertical Z direction Piezo to allow this nanopipette to hop over the surface of platelets. A 0.3–0.4% reduction of the ion-current flowing into nanopipette was set to maintain a constant separation between the nanopipette and platelet surface. During scan, the nanopipette performed pre-scan hopping distance at 3000 nm and 400 nm amplitude for high-resolution scanning. The nanopipette falling rate was set to 50 nm/ms, meanwhile its rising rate was increased to 500 nm/ms. An adaptive imaging protocol of HPICM was used for scanning platelets [22]. Briefly, HPICM pre-scanned a designated area to estimate the overall roughness by measuring the difference in heights at the four corners. If a platelet was detected in the area, HPICM subsequently scanned this platelet at a higher resolution of 256 256 pixels. In contrast, HPICM scanned a flat surface identified in the pre-scan at a lower resolution of 128 128 pixels. The operation time of this adaptive scanning was further reduced by obtaining high-resolution scan immediately after the pre-scan at the four corner points. This is particularly useful for
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imaging living platelets because it ensures that platelets undergo minimal shape changes between pre-scan and high-resolution scan. This adaptive operation protocol also accelerated the speed of HPICM scan without the loss of high-resolutions. At this setting, each scan took approximately 6–9 min. During the entire scan period, platelets were bathed in L15 medium and maintained at 24–28 C. Acquired raw topography data were bilinearinterpolated using ScanIC Image Viewer version 1.0 (Ionscope Ltd, Royston, UK) to produce final images of 512 512 pixels. Platelet imaging by AFM For comparison, human platelets were scanned by AFM. AFM was performed in liquid at a rate of 0.3 Hz in a contact mode with DNP-S cantilevers (fo ¼ 12–24 kHz, k ¼ 0.06 N/m, Bruker Corporation, Santa Barbara, CA) using a BioScope IITM Controller (Bruker Corporation) integrated into a Nikon TE2000-E inverted optical microscope (Nikon Instruments, Inc. Lewisville, TX) as previously described [27]. The graphic program Research NanoScope version 7.3 was used to convert digital data into topographical images. The time required to scan a 50 50 mm2 area with a typical resolution of 512 512 pixels by AFM was approximately 26 min. Platelet imaging by SEM For comparison, human platelets were also scanned with SEM. Carbon grids were coated with fibrinogen over night at 4 C and then incubated with PRP for 30 minutes at 37 C. After nonadherent platelets were removed by washing, platelets on grids were fixed with 3% of glutaraldehyde, dehydrated with increasing concentrations of ethanol (70%, 90% and 100%), and rehydrated with tert-butyl alcohol as previously discussed [27]. SEM micrographs were recorded with JSM-6380Lv SEM (JEOL Ltd., Tokyo, Japan). Statistical analysis HPICM images were processed and analyzed by ScanIC Image Viewer version 1.0 program (Ionscope Ltd, UK). All data were analyzed using Origin 8.0 program (Originlab Corporation, Northampton, MA). Quantitative values were presented as mean ± SEM. All data were analyzed by pair comparison. A difference between means at the level of p50.05 was considered statistically significant; p50.005 was statistical highly significant.
Results Continuous HPICM observations of live platelets With technical settings discussed in the ‘‘Method’’ section, we were able to detect live platelets adherent to fibrinogen at a spatial resolution of 30 nm, without direct contacts between platelets and the nanopipette. Platelets remained adherent and stable over a 60–90 minutes period of repeated scans (6–10 cycles) by HPICM [27]. Figure 1 shows HPICM topographical images of platelets adherent to fibrinogen taken every 9 minutes. Without exposure to an agonist, platelets adherent to fibrinogen presented in two distinct morphologies: low-density spread shape (LDSS) and high-density bubble shape platelets (HDBS) (note morphological changes of six platelets highlighted by white dotted ellipses). This difference in density indicates the different heights of adherent LDSS (0.51 ± 0.13 mm, n ¼ 35) and HDBS (1.88 ± 0.54 mm, n ¼ 36) platelets. In addition, LDSS platelets were larger (5.68 ± 1.44 mm, n ¼ 56) than HDBS platelets (2.95 ± 0.62 mm, n ¼ 72), suggesting that LDSS platelets spread completely on fibrinogen. Some HDBS platelets gradually
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DOI: 10.3109/09537104.2014.940888
Morphology imaging of live platelet by HPICM
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Figure 1. Repetitive HPICM scans of live platelets on fibrinogen substrate. Six images were acquired every 9 minutes over a 45 minutes time-lapse scanning of a selected 50 50 mm area.
Figure 2. High-resolution HPICM images of changed morphology from HDBS to LDSS. (A) Three images constituted a sequence of 8 minutes time-lapse scanning. (B) Scanning profile changes of the solid line via pseudo-nuclear center in A from 0 minutes (thick black line trace) to 8 minutes (grey line trace) to 16 minutes (thin black line trace).
became a LDSS morphology (Figure 1, arrow), while others remained unchanged, but with slightly shifted their positions (white solid ellipse) during repeated scans. In contrast, LDSS platelets underwent little change in morphology during the same period. These data suggest that the membranes of live platelets are highly dynamic and the two distinct morphologies of LDSS and HDBS likely represent two interchangeable stages of platelet adhesion to and spreading on fibrinogen. One set of high-resolution time-lapsed HPICM scans detected this gradual change in morphology from HDBS to LDSS in approximately 16 minutes (Figure 2). During spreading, the height of HDBS platelets at the pseudo-nuclear center reduced from 1.74 ± 0.50 mm to 0.51 ± 0.18 and its diameter increased from 2.66 ± 0.41 mm to 5.33 ± 1.10 mm, leading to an increase in the surface area from 14.66 ± 4.99 mm2 to 28.70 ± 10.95 mm2 without significant change in volume (n ¼ 11). A scanning profile of the platelet cross-section marked by the straight-line across through three time-lapse frames show such morphological changes during the scanning period (Figure 2B). These data for the first time showed platelet membrane fluidity and spreading in high resolution. Because of its ability to repeatedly scan live platelets with no direct light and heat exposure, HPICM is ideally to observe the dynamic process of platelet spreading on different matrixes. Platelets were allowed to adhere to either human fibrinogen, which is mediated by the platelet integrin aIIbb3 [28,29], or
positively charged poly-D-lysine, which mediated platelet adhesion through an electrostatic interaction [30]. Both HDBS and LDSS morphologies were also observed from those adherent platelets on poly-D-lysine (Figure 3E–H). However, platelets on poly-D-lysine were significantly larger and thicker (with a big volume) than those on fibrinogen. Both pseudo-nuclear center height and cell volume of HDBS platelets on poly-D-lysine were significantly larger than those on human fibrinogen (Table I). Morphology of platelets observed by SEM and AFM We compared morphological characteristics of adherent platelets on fibrinogen observed by HPICM to those detected by AFM and SEM (Figure 4). An AFM cantilever probes platelets by touching their surfaces and was able to detected HDBS and LDSS morphologies of adherent platelets [27]. However, an AFM cantilever removed a substantial number of adherent platelets (35%) during repeat scans (data not shown), generating different ratios of HDBS to LDSS platelets on fibrinogen (0.21 ± 0.03 for AFM, n ¼ 6, vs. 0.94 ± 0.54 for HPICM, n ¼ 10). This observation suggests that the mechanical force loading on platelets by the cantilever may have pulled platelets from the substrate and HDBS platelets were more sensitive to this mechanical pulling force. To address this concern, we also scanned platelets fixed by paraformaldehyde (Figure 4B). The fixation significantly reduced the number of platelets that were removed by the cantilever, but
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Figure 3. HPICM images of HDBS and LDSS morphologies of live human platelets adherent to fibrinogen (A–D) and poly-D-lysine (E–H). B–D and F–H are gradually zoom-in scans of HDBS and LDSS platelets corresponding to those white-dotted-squares marked areas of B–D and F–H in figures A–B and E–F, respectively. Table I. Measurements of platelets adherent to fibrinogen and poly-D-lysine.
HDBS (n ¼ 36) Height (mm) Volume (mm3) LDSS (n ¼ 35) Height at peudo-center (mm) Volume (mm3)
Fibrinogen
Poly-D-lysin
p Value
1.88 ± 0.54 11.00 ± 3.75 0.51 ± 0.13 13.44 ± 4.47
2.63 ± 0.65 17.02 ± 5.90 0.89 ± 0.45 18.23 ± 11.57
50.005 50.005 50.005 50.05
Figure 4. Platelets adherent to fibrinogen-coated surface imaged by SEM (A) and AFM (B). Arrows pointed platelets had low density spread shape (LDSS) and circle surrounded platelets had high density bubble shape (HDBS).
some of pseudo-center of HDBS platelets were scratched by the cantilever because the distance between the cantilever and platelet is fixed. Platelets adherent to fibrinogen were also fixed and imaged by SEM (Figure 4A). Both HDBS and LDSS platelets were identified at a ratio similar to that of HPICM. However, both AFM and SEM were unsuitable to detect dynamic changes in living platelet morphologies.
Discussion We have shown that HPICM is an effective tool in monitoring morphological changes of adherent platelets in real-time. Using this HPICM technology, we identified two distinct morphologies of live platelets adherent to fibrinogen: fully spread (LDSS) and
non-spreading (HDBS). These morphologies are highly dynamic in real-time and HDBS platelets gradually became LDSS morphology in 10–20 minutes. These two morphologies were also observed from platelets adhered to poly-D-lysine, suggesting that their formation is independent of the interaction between fibrinogen and the platelet integrin aIIbb3. However, this morphological distinct is important because we have recently shown that HDBS, but not LDSS, platelets are very sensitive to agonist-induced microvesiculation [27]. These dynamic and heterogeneous characteristics of platelets are distinct from neurons and epithelial cells that have previously been studied with this technology [22–25]. A key technical feature of HPICM is its hopping probe, which provides a sensitive tip-sample separation control to perform non-contact surface profiling [21–25] and will not disturb the
Morphology imaging of live platelet by HPICM
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DOI: 10.3109/09537104.2014.940888
platelet membrane. In contrast, a conventional AFM requires a suitable surface viscosity and stiffness of a cell. Because the distance between a surface and a cantilever is fixed, AFM is less flexible in probing a cell that undergoes morphological changes. For example, a setting for tall HDBS platelets may sacrifice the resolution for LDSS platelets, whereas the tip of AFM cantilever could scratch the membrane surface at the top of a HDBS platelet when the distance was set for fully spread platelets, resulting in image distortion as seen in Figure 4(B) (highlighted by white dotted circle). Another drawback for AFM imaging is that an acquisition time needs to be increased when a dynamic process is to be visualized. High-speed AFM has been recently used to directly visualize the dynamics structure and processes of biological molecules in physiological solutions [31–33], but its application in live cells is few. This technical obstacle of a longer acquisition time is also addressed by HPICM, which uses an adaptive resolution scanning protocol to reduce the acquisition time for one frame of scanning of living platelets in a 50 50 mm2 area to 6–9 minutes. In conclusion, upon binding to the subendothelial matrix, platelets are activated and undergo drastic shape changes that are critical for platelet adhesion and aggregation [34–36]. These morphological changes of platelets have been widely reported, but are mostly observed after platelets were fixed with a chemical fixative. HPICM overcomes several key technical difficulties associated with confocal microscopy, AFM and SEM to offer high-resolution monitoring morphological changes and defining their physiological significance in real-time without inducing mechanical and chemical damages to platelets. Time lapse HPICM scans offer a unique opportunity to monitor the agonist-induced dynamic changes in platelet morphologies that have been difficult to detect with other imaging technologies [27]. HPICM can also be combined with patch-clamping technology using the same nanopipette [24, 25], so that one can simultaneously monitor platelet shape changes and the activity of ion channels during platelet activation, which has not been possible with current imaging technologies.
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Declaration of interest This work was supported by Tianjin Science and Technology Support Program of China (No.14ZCZDSY00020), Tianjin Natural Science Foundation of China (No.13JCYBJC21900 and 12JCYBJC31500), National Natural Science Foundation of China (No.31300828, 81330029, and 81271361), and the National Heart, Lung, and Blood Institute, NIH, USA (grants HL71895 and HL85769).
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