Special thanks to Dr. Clare M. Waterman for mentorship and discussions, Michelle Baird and Mike Davidson for providing many of the fluorescent cDNA constructs used in these studies, and Kathryn Applegate and Gaudenz Danuser for developing plusTipTracker MT analysis software. This work was supported, in part, by a grant to K.A.M from the National Heart Lung and Blood Institute (NHLBI) and the National Institutes of Health (NIH). (Grant: 4K22HL113069).

The authors declare that they have no competing financial interests.

Using HUVECs in Morphology and Migration Experiments. 
Live-cell imaging of EB3-labeled MT growth dynamics within HUVECs requires appropriate and reproducible cell culture conditions in order to measure and evaluate changes in MT growth and HUVEC morphology, such that they can then be assigned to an ECM signaling cue. HUVECs represent an excellent model for studying cell shape and motility. They are highly amenable to transfection with fluorescently-labeled cDNAs, they exhibit dramatic morphologies in response to both soluble and physical signaling cues, and they maintain the ability to organize themselves into vascular tubes when provided appropriate cell culture conditions65-70. Primary cell cultures, such as HUVECs, require careful handling and monitoring of cell passage number in order to ensure the cells are healthy and that they will behave normally during experimentation. For example, when conducting experiments investigating the cytoskeleton and/or cell morphology, HUVECs should not be used for experiments beyond the tenth cell culture passage, as HUVECs typically begin to display non-uniform morphologies around passage twelve to fifteen.
Cell density is also a critical regulator of HUVEC health and proliferation. HUVEC branching investigations (see Protocol 10.1-10.4, above) use relatively low cell density cultures in order to provide the cells physical space to interact with their ECM and to extend branched protrusions. In contrast, in wound-edge migration studies (see Protocol 10.5-10.8), HUVECs are culture in a monolayer prior to wounding. As such, wound-edge HUVECs display a relatively un-branched morphology, extend leading edge protrusions, and exhibit cell-cell interactions with their immediate neighbors as they migrate into the wound.
Caveats and Advantages of Tracking MT Dynamics in Living HUVECs. 
Spinning disk microscopy is an excellent approach to test experimental hypotheses that require confocal image clarity with high time-resolution. With the appropriate camera and laser module, this microscopy approach is sensitive to low emission fluorescence and can aid in reduced photobleaching compared to other types of fluorescence microscopy. Importantly, this approach is also well suited for moderate to high time-resolution imaging as is required for generating comprehensive measurements of MT growth dynamics using the EB3 labeling approach, in a variety of different cell types.
While the EB3 methodology of labeling growing MT plus-ends has distinct advantages over other methods of fluorescently labeling MTs, it also has unique disadvantages. For example, there is a proportion of the MT array that is not actively growing at any given time, including both the population of disassembling MTs and the population of stabile, non-dynamic MTs71-74. EB3 would not label these two populations of MTs and therefore the contribution of these two populations would not be included in experimental data analysis. Alternative methodologies for evaluating the entire MT array have focused on expression of fluorescently-labeled tubulin, which incorporates into all populations of MTs and allows for identification of growing, shrinking, and stabile MTs. However, because of the relatively high density of the MT array near the center of the cell and adjacent to the microtubule organizing center (MTOC), labeling the entire MT array creates a disadvantage in performing image analysis of MT ends that are not near the cell periphery75-77. Experiments using two-color co-labeling of MTs with fluorescent tubulin and fluorescent EB3 in live cells have revealed, qualitatively, that the vast majority of tubulin-labeled MTs are also EB3-labeled78. Additionally, there has not been an efficient method of automated tracking of MTs labeled with fluorescent tubulin. This means that data collection and measurements of MT dynamics in cells expressing fluorescent tubulin require hand tracking of individual MTs, a process that is error prone and tedious. As a result, automated computational analysis of EB3 labeled MTs has become the preferred methodology for measuring MT dynamics as it can be applied to MT dynamics experiments across numerous cell types and within a variety of experimental regimes35,79.
Linking MT Growth Dynamics to HUVEC Morphology and Migration. 
From the perspective of a cytoskeletal investigator, one extremely useful adaptation of the plusTipTracker automated tracking analysis is the ability to measure and evaluate both whole cell and regional changes in MT growth dynamics. The requirement for a cell to become polarized is an essential pre-requisite for directional cell migration. As such, polarized signaling cues have long been known as key driving factors for directional cell migration30,31,51,54,56,80,81. For example, soluble signaling molecules such as VEGF induce actin polymerization and MT growth toward the VEGF cue in endothelial cell types2,82-87. Additionally, in response to physical interactions with the ECM (for example, compliance and dimensionality mechanosensing), HUVECs modulate their MT dynamics by locally regulating MAPs to promote MT growth within elongating branched protrusions, and this serves to guide HUVEC migration in the direction of the cue5,31,88. Thus, polarization in response to extracellular signaling cues highlights the need for experimental evaluation of localized changes in cytoskeletal dynamics.
Simultaneous live-cell imaging of MT growth dynamics (using the EB3 approach) and HUVEC cell migration is extremely difficult to perform because MTs grow on a time scale of seconds, while changes in cell morphology and directional migration occur on a time scale of hours. Yet both processes are intimately linked. Moreover, continuous, rapid imaging of fluorescently-labeled EB3 on the second time scale leads to photobleaching of the EB3 signal. Attempts to overcome this issue by conducting short imaging series of EB3 (1-2 min acquisition series) every 30 minutes have been successful, but could only be accomplished in a small number of ECs that had optimal expression of EB3 and that did not display adverse effects to repeated laser illumination5.
An alternative approach that allows for interpretations of how MT growth dynamics contribute to changes in cell morphology and migration is the plusTipTracker Region of Interest (ROI) tool35 (see Protocol 6.6). This methodology allows for whole cell MT growth tracks to be sub-divided into user-defined regions, from which MT growth tracks can then be extracted and analyzed. For example, comparisons of &#34;branched&#34; versus &#34;non-branched&#34; regions of the cell have revealed that MTs grow more slowly and more persistently within branched regions as compared to the non-branched cell periphery5. In polarized wound-edge HUVECs, comparisons of the leading edge and trailing edge MT growth dynamics revealed that MCAK-induced MT disassembly is inhibited specifically within the leading edge, but not the trailing edge. Regional evaluation of leading and trailing edge MT growth dynamics in HUVECs expressing Rac1 and/or phosphorylation mutants of MCAK further revealed that Rac1-induced activation of Aurora-A kinase specifically and locally inhibits MCAK activity within the leading edge to drive HUVEC polarization and directional migration31. Thus, the combination of automated tracking of MT growth dynamics and regional evaluation of MT growth dynamics within the branched protrusions and/or the leading edge of wound-edge HUVECs has allowed us to link MT growth dynamics to HUVEC morphology and migration.
The protocols described are designed to provide a generally straightforward and highly repeatable methodology for HUVEC culture and experimental investigation. Nevertheless, many of the protocol details are complex and time consuming, which, in turn, provides opportunity for errors or difficulties in conducting the experimental methodology. While it was attempted to address potential problems throughout the protocol, here is provided additional, detailed troubleshooting of potential problems that are less common or otherwise abbreviated as notes within the methodology protocols.
Related to coating coverslips with polyacrylamide substrates (Protocol 2.2), a common problem that arises is the complex nature of activating glass coverslips, coupling polyacrylamide to the glass, activating polyacrylamide, and then coupling ECM to the polyacrylamide. One particularly difficult and potentially problematic step is culturing HUVECs onto the fibronectin/polyacrylamide gel following exposure of the polyacrylamide-coated coverslips to 2 mM sulfo-SANPAH (step 2.2.2.6, above). It is critical to the success of culturing HUVECs on these substrates that sulfo-SANPAH be completely removed from the experimental system following ECM conjugation. This can be best accomplished by washing with ddH2O and incubating the coverslips in ddH2O on a shaker overnight. If cells cultured on the polyacrylamide ECMs do not survive, or if viability is less than expected, increased and repeated washing of sulfo-SANPAH after conjugation to fibronectin (step 2.2.2.9) is recommended to potentially alleviate this problem.
Related to Protocol 3 (cDNA transfection and HUVEC culture on a coverslip), a major influence to the success of the electroporation procedure is the handling of the HUVECs both during trypsinization of the cells to remove them from the plastic flask, and following the conclusion of the electroporation (steps 3.3-3.4, above). It is critical to carefully monitor the HUVECs while in trypsin, and not to exceed the time needed for the cells to &#34;round-up&#34; on the surface of the flask (typically 1-2 min). Excessive time in trypsin is a major cause of HUVEC death, which is typically not realized until the cells are plated onto fibronectin-coated coverslips (step 3.4) and then fail to attach to the substrate.
Of similar importance, HUVECs (and most primary cell types) do not fare well when suspended for long times in the electroporation buffer. During larger-scale experiments, for example, where the user has five or more conditions that require electroporation, it is better to keep the cells pelleted in individual tubes (1 tube per transfection) and to mix them, one-at-a-time, in the electroporation buffer immediately before electroporation. This approach minimizes incubation time in electroporation buffer and maximized HUVEC survival. Additionally, immediate rescue in complete culture media is also critical following electroporation. Rescue delays of one minute or more following electroporation can result in near complete HUVEC death and thus should be avoided.
Related cell migration assay (Protocol 9), the technique of culturing HUVECs at extremely high density depends on a critical modification (described in step 9.2) of the traditional HUVEC cell culture methodology (described in Protocol 3) that requires drying of the fibronectin coated coverslip in order to enable HUVEC culture into a very small area of the coverslip. Often times, if the coverslip is not completely dried or if the prior coating of the coverslip with fibronectin (steps 2.1 or 2.2) was not efficient, the media-containing cells that is placed on the coverslip will lose its surface tension and expand to the edges of the coverslip, thereby reducing the density of cells on the coverslip. One method to troubleshoot this potential problem is to aspirate the media from the coverslip and then place the coverslip (still in its 35mm dish) into the back of the biosafety cabinet for 5-10 min. This adaptation can help to allow the flow of air within the cabinet to assist in drying the dish. If any visible liquid remains on the coverslip after air drying, aspirate the liquid spots and again allow to air dry for 5-10 additional min in the biosafety cabinet. It is important to note that there is no way to completely avoid this potential problem, but it does become less of an issue with repeated practice of the protocol. It is also a troubleshooting recommendation that when preparing coverslips for the cell migration assay (or any assay, in general), to prepare extra coverslips and have extra HUVECs ready-to-go in case of problems along the way.
With these troubleshooting considerations in mind, it is also important to highlight the utility of the protocols discussed and their implications in future cell biology research. The applicability to the polyacrylamide approach is wide-ranging, and includes not only two-dimensional studies of cell-ECM engagement (described above), but also three-dimensional investigations5. This application is critical to increasing our understanding of how HUVECs regulate cell shape and migration in physiological environments and should provide investigators a basic understanding of how morphological changes are coordinated with cytoskeletal changes. While the protocols described here are focused on measurements of MT growth dynamics, the majority of the protocols can and should be adapted to measure any number of other microscopically measureable aspects of cell-ECM interactions. With respect to MT dynamics, there are numerous MAPs, including at least 14 families of kinesins89, each with a potential role in contributing to HUVEC polarity and directed migration, and all of which can be investigated using the protocols presented.
Future investigations should also be targeted at cell-ECM engagement in normal versus diseased states. HUVEC protocols presented here are applicable to investigations of normal vascular homeostatic processes, as well as to disease states including heart disease, retinopathy, tumor growth and metastasis, to name a few. Conjugating visibly transparent ECMs and polyacrylamide substrates of amenable compliance with microscopic studies will allow for cell-ECM engagement studies such that endothelial cell vascular angiogenesis can be monitored with high spatial and temporal resolution. These types of experimental approaches have already begun to reveal important differences in endothelial cell shape, polarity, migration, and the effects of proteins that regulate these processes5,31,90. Future efforts should aim to identify how the combination of ECM compliance and dimensionality mechanosensing may serve to locally control proteins that target and regulate cytoskeletal dynamics.

Angiogenesis is triggered by extracellular signaling cues that promote endothelial cells (ECs) to polarize, migrate with directional specificity, and form new vasculature in tissues that require blood supply. Contributing factors thought to drive angiogenesis are signals from the extracellular matrix (ECM). In response to physical cues, ECs extend new branches with directional specificity to guide directional movement in formation of the vascular network1,2. The architecture of EC morphology and branching is defined by localized regulation of the microtubule (MT) cytoskeleton in a way that is coordinated with the regulation of acto-myosin contractility3. In order for EC branches to form, myosin-II must be locally downregulated, resulting in localized membrane protrusions4. At the same time and in the same location within the cell, MT growth dynamics become modified resulting in slow and persistent MT growth to promote branch elongation and stability5. This coordinated cytoskeletal regulation allows ECs to polarize their morphology to facilitate directional migration.
The development of a polarized cell morphology requires polarized MT assembly6. In migrating epithelial cells, MTs grow slowly and persistently specifically at the leading edge of the cell7-9; while in neurons, the initiation and elongation of axons or dendrites requires that MTs are not only present, but that they are dynamically undergoing the processes of assembly and disassembly10-15. In this way, localized control of MT growth dynamics determines the architecture of the cell and allows for rapid remodeling of cell morphology as ECs navigate through their ECM.
MT growth dynamics are regulated by families of molecular motor proteins and non-motor proteins, termed Microtubule Associated Proteins or Microtubule Interacting Proteins (henceforth referred to as MAPs). MAPs can be locally activated or inactivated, typically through signaling cascades initiated at the cell-ECM interface16,17. Once active, MAPs function by binding to the MT and altering the kinetics of MT assembly and disassembly; thereby functioning to locally regulate MT dynamics in order to promote cell shape and polarity18-20. Mitotic Centromere Associated Kinesin (MCAK) is a Kinesin-13 family MAP that binds to MT ends and couples ATP hydrolysis to MT disassembly21-23. This functional role of MCAK is known to be critical within the mitotic spindle24-28, as well as during interphase29,30. Within interphase ECs, MCAK-mediated MT disassembly is regulated by localized Aurora-A induced phosphorylation of MCAK in response to wound-edge induced activation of the Rac1 small GTPase. The outcome of this signaling cascade is the inhibition of MCAK at the leading edge of ECs, resulting in polarized MT growth toward the leading edge and MCAK-induced MT disassembly at the trailing edge of migrating ECs31.
Traditional methodologies for measuring MT growth dynamics have typically used hand tracking of either fluorescently-labeled tubulin or, more recently, fluorescently-labeled MT End Binding protein 1 or 3 (EB1 or EB3, respectively). The fluorescent tubulin approach has the advantage of labeling the entire MT array. However, because the majority of mitotic cell types maintain a radial array of MTs during interphase8,32,33, MTs near the centrosome are very dense, and as a result, tracking MT growth and disassembly with fluorescent tubulin is usually limited to the peripheral regions of the cell. Because of these limitations, utilization of fluorescently-labeled EB proteins (EB1 or EB3) has been the more common approach to measuring MT dynamics. Because EBs only label the growing plus-ends of MTs, they appear as small (1-3 &#956;m), brightly fluorescent comets, thus providing an easily discernable marker of MT polymerization with very limited background fluorescence34-37. While the fact that EBs label only a subset of the MT array represents a major strength of the EB approach, it also highlights one important limitation, that non-growing MTs are not labeled by the EB3 approach. Nevertheless, the ability to measure and track EB3 comets over time and to visualize MT growth within sub-cellular regions make this approach ideal for applications that require regional evaluation of MT organization.
Another critical limitation of traditional methodologies for measuring MT growth dynamics has been very large data sets and the time required for data analysis. This limitation stems from the need for hand tracking of hundreds of EB comets at high time resolution. Moreover, these measurements need to be done in a statistically significant number of cells and also performed under a variety of control and experimental conditions. Recently, these constraints have been overcome through computational analysis techniques specifically directed at tracking EB3 comets using time-lapse microscopy in living cells35. When used appropriately, this computational approach serves to both restrict the potential for human error associated with hand-tracking in addition to providing for a high-throughput method of measuring MT polymerization. Computational analysis of MT dynamics using the MatLab-based software package, plusTipTracker, allows for measurements of MT growth by tracking fluorescently-labeled EB3 comets5,31,35,38. Additionally, plusTipTracker software allows for calculations of MT catastrophe events (a.k.a. disassembly) based on the disappearance of EB3 comets within a growing MT track, and allows for sub-cellular (a.k.a. regional) analysis of MT growth dynamics within user-defined regions of interest. For our studies, MT growth dynamics were compared within branched regions versus non-branched regions, or within leading versus trailing cell edges. Data output from plusTipTracker software can also be used to generate color-coded track overlays for individual cells and sub-cellular regions.
Here we describe the methodology used to investigate the role of MCAK in modulating MT growth dynamics and the contribution of this MT depolymerase to the regulation of EC branching morphology and directional migration. Our approach used a primary human cell line, Human Umbilical Vein Endothelial Cells (HUVECs), which are easily cultured and highly amenable to electroporation-induced, transient transfection of plasmid cDNA. We then used high-resolution, time-lapse fluorescence microscopy of HUVECs transfected with MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics transfected HUVECs. PlusTipTracker-based computational image analysis of EB3-labeled MT plus ends was to measure MT growth speeds and MT growth lifetimes in time-lapse images, to measure and compare effects of experimental manipulations on MT growth dynamics. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration. These techniques are applicable to investigations of any MAP that can be fluorescently labeled and expressed in live cells.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Aurora A Inhibitor I</td>
			<td>Selleck Chemicals</td>
			<td>S1451</td>
			<td></td>
		</tr>
		<tr>
			<td>2% Bis solution</td>
			<td>Bio-Rad</td>
			<td>1610142</td>
			<td></td>
		</tr>
		<tr>
			<td>3-aminopropyltrimethyloxysilane</td>
			<td>Sigma-Aldrich</td>
			<td>A3648</td>
			<td></td>
		</tr>
		<tr>
			<td>40% acrylamide</td>
			<td>Bio-Rad</td>
			<td>1610140</td>
			<td></td>
		</tr>
		<tr>
			<td>405, 488, 561, 640 nm high power MLC400 laser combiner module</td>
			<td>Agilent Technologies</td>
			<td>99462</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium persulfate</td>
			<td>Sigma-Aldrich</td>
			<td>A3678</td>
			<td></td>
		</tr>
		<tr>
			<td>anti&#8211;mouse IgG immunoprecipitation beads</td>
			<td>TrueBlot Reagents; Rockland Inc.</td>
			<td>88-1688-31</td>
			<td></td>
		</tr>
		<tr>
			<td>anti&#8211;rabbit IgG immunoprecipitation beads</td>
			<td>TrueBlot Reagents; Rockland Inc.</td>
			<td>88-7788-31</td>
			<td></td>
		</tr>
		<tr>
			<td>ET455/50m, DAPI-ET Emission Filter</td>
			<td>Chroma Technology Corp.</td>
			<td>ET455/50m</td>
			<td></td>
		</tr>
		<tr>
			<td>ET525/36m Emission Filter</td>
			<td>Chroma Technology Corp.</td>
			<td>ET525/36m</td>
			<td></td>
		</tr>
		<tr>
			<td>ET605/70m Emission Filter</td>
			<td>Chroma Technology Corp.</td>
			<td>ET605/70m</td>
			<td></td>
		</tr>
		<tr>
			<td>ET700/75m Emission Filter</td>
			<td>Chroma Technology Corp.</td>
			<td>ET700/75m</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin Fraction V</td>
			<td>RPI Products</td>
			<td>A30075-100.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Clara cooled charge-coupled device camera</td>
			<td>Andor Technology</td>
			<td>77026010</td>
			<td></td>
		</tr>
		<tr>
			<td>Cy3 donkey anti-rabbit</td>
			<td>Jackson ImmunoResearch</td>
			<td>711-165-152</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Thermo Fisher Scientific</td>
			<td>14190250</td>
			<td></td>
		</tr>
		<tr>
			<td>Dyelight 649 goat anti-rat</td>
			<td>Jackson ImmunoResearch</td>
			<td>112-496-068</td>
			<td></td>
		</tr>
		<tr>
			<td>EGM-MV SingleQuot Supply &#38; Growth Factors (contents shown below)</td>
			<td>Lonza</td>
			<td>CC-4143</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum 25 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine brain extract 2.0 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamycin sulfate 0.5 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Epidermal growth factor 0.5 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrocortisone 0.5 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin (10,000 IU Pen/mL, 10,000 ug Strep/mL)&#160; 10.0 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma-Aldrich</td>
			<td>E3884-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Electronic shutter</td>
			<td>Sutter instrument</td>
			<td>77016099</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>EMD Millipore</td>
			<td>FC010</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Thermo Fisher Scientific</td>
			<td>S80026</td>
			<td></td>
		</tr>
		<tr>
			<td>HALT Protease inhibitor</td>
			<td>Thermo Fisher Scientific</td>
			<td>PI-78425</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>HRP goat anti-mouse</td>
			<td>Jackson ImmunoResearch</td>
			<td>115-036-062</td>
			<td></td>
		</tr>
		<tr>
			<td>HRP goat anti-rabbit</td>
			<td>Jackson ImmunoResearch</td>
			<td>111-036-045</td>
			<td></td>
		</tr>
		<tr>
			<td>Immuno-Blot PVDF Membrane</td>
			<td>Bio-Rad</td>
			<td>162-0174</td>
			<td></td>
		</tr>
		<tr>
			<td>Lanolin</td>
			<td>Thermo Fisher Scientific</td>
			<td>8006-54-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>M2643-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon TiE</td>
			<td>MEA53100</td>
			<td></td>
		</tr>
		<tr>
			<td>Motorized stage</td>
			<td>ASI Technologies</td>
			<td>MS-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Mounting medium</td>
			<td>Dako</td>
			<td>CS70330-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-Aurora A</td>
			<td>Abcam</td>
			<td>ab130156</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-GAPDH</td>
			<td>Abcam</td>
			<td>ab9484</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-MCAK</td>
			<td>Abcam</td>
			<td>ab50778</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-tubulin (DM1A)</td>
			<td>Cell Signaling Technology</td>
			<td>3873S</td>
			<td></td>
		</tr>
		<tr>
			<td>Nail Polish</td>
			<td>Electron Microscopy Sciences</td>
			<td>72180</td>
			<td></td>
		</tr>
		<tr>
			<td>NIS Elements software</td>
			<td>Nikon</td>
			<td>MQS310000</td>
			<td></td>
		</tr>
		<tr>
			<td>Nucleofector Kit V</td>
			<td>Lonza</td>
			<td>VACA-1003</td>
			<td>Contains the electroporation buffer</td>
		</tr>
		<tr>
			<td>Paraffin</td>
			<td>Thermo Fisher Scientific</td>
			<td>P31-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000IU Pen/mL,10,000ug Strep/mL)</td>
			<td>MP Biomedicals</td>
			<td>91670249</td>
			<td></td>
		</tr>
		<tr>
			<td>Petroleum jelly</td>
			<td>Dow Corning</td>
			<td>2021854-1113</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipes</td>
			<td>Sigma-Aldrich</td>
			<td>P6757-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>plusTipTracker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-GFP</td>
			<td>Abcam</td>
			<td>ab290</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti&#8211;pT288&#8211;Aurora A</td>
			<td>Abcam</td>
			<td>ab18318</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti&#8211;&#945;-tubulin</td>
			<td>AbD Serotec</td>
			<td>MCA78G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium borohydride</td>
			<td>Thermo Fisher Scientific</td>
			<td>678-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinning disk&#160;</td>
			<td>Andor Technology</td>
			<td>Yokagawa CSU-X1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sulfo-SANPAH</td>
			<td>Thermo Fisher Scientific</td>
			<td>PI-22589</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperSignal West Pico Chemiluminescent Substrate</td>
			<td>Thermo Fisher Scientific</td>
			<td>34080</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Thermo Fisher Scientific</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Corning</td>
			<td>MT25053CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Thermo Fisher Scientific</td>
			<td>BP337</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-resolution time-lapse imaging and automated analysis of microtubule dynamics in living human umbilical vein endothelial cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

HUVEC culture for live-cell microscopy 
Live-cell imaging of HUVECs expressing fluorescent proteins requires that HUVECs are maintained in an environment that is suitable for normal cell growth and maintenance, as well as normal protein expression and function. Figure 1 shows a schematic depiction of the protocol used to prepare a buffered and closed system that maintains optimized optical characteristics for collecting time-lapse images of live cells. Thin strips of double-sided tape are placed onto a glass slide (Figure 1A) and then the coverslip containing the cultured HUVECs is inverted on top of the tape such that the cells are facing the slide (Figure 1B). In this way, the tape provides a small gap between the two glass surfaces, and creates a space in which the cells can be bathed in appropriate cell culture media (Figure 1C; see Protocol 4, above). In a final step, the glass coverslip is sealed to the glass slide using a petroleum jelly: lanolin: paraffin wax (1:1:1 mixture, VALAP; Figure 1D) to establish a closed system. This cell culture approach was used for both the shorter time sequence imaging of EB3 comets and for the longer time sequence branching and migration experiments. Additionally, the wax-like consistency of VALAP allows for easy removal from the glass slide and coverslip at the end of an imaging experiment such that supplemental approaches including fixation and immunolabelling (see Protocol 8), can be performed on the same coverslip and cell sample that was visualized using the live-cell imaging approach.
<img alt="Figure 1" src="/files/ftp_upload/54265/54265fig1.jpg" /> 
Figure 1: Methodology of HUVEC culture for live-cell microscopic imaging. (A) Cut 2 pieces of double-sided tape into 0.65 cm x 2.50 cm rectangular strips and secure one piece of double-sided tape horizontally along the top edge of the slide, and the other piece, along the bottom edge of the slide. It is important to allow a small amount of space between the tape and the slide edge for sealing the chamber as described in (D). (B) Using forceps remove the coverslip containing HUVECs and invert the coverslip (cell side down) on top of the pieces of tape. (C) Using a micropipette, add 40 &#956;l of imaging media (complete endothelial basal medium supplemented with 25 &#956;M HEPES) between the coverslip and the glass slide. Ensure that the space between the coverslip and slide is completely filled with imaging media and avoid bubbles. Use an aspirator along the edge of the coverslip to remove excess media, if necessary. (D) Use warm VALAP (1:1:1 vaseline : lanolin : paraffin) to seal around the edges of the coverslip. Check for imaging media leakages by pushing around gently on the coverslip. Use additional VALAP to remedy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Automated tracking of fluorescently-labeled EB3 allows for comprehensive analysis of whole cell and regional MT growth dynamics. The collection of clear, distinct, and highly time and space-resolved microscopic images of the EB3 protein is essential to the analysis of EB3 movements within the cell. HUVEC expression of fluorescently-labeled EB3 varies widely from cell to cell and fluorescence intensity typically increases within the same cell over a period of hr. Thus, it is important to consider and monitor these changes in expression profile in order to identify potential effects of increased EB expression on MT growth dynamics. This is best accomplished by evaluating grey-scale fluorescence intensity profiles for each imaged cell, and then restricting data analysis to those cells that display a defined range of grey-scale expression profiles. Initial evaluation of expression profile data sets is critical, and should be compared with MT growth dynamics data that is obtained by plusTipTracker software for each cell. Expression profiles can be plotted against MT growth speed and MT growth lifetime for each cell within an experimental group, in order to determine the desired range of grey-scale expression and to identify potential outliers.
Figure 2 highlights a HUVEC cultured on a fibronectin substrate (see Protocol 2.1, above) and expressing near optimal levels of mApple-labeled EB3. Fluorescent EB3 comets are shown in a time-lapse series of images over a period of 12 sec (Figure 2A). Analysis of EB3 comets using plusTipTracker software involves an automated, frame-by-frame EB3 comet detection (green and yellow dots, Figure 2B). Comets detected within each frame are then linked, dependent upon their change in position from frame-to-frame, to generate EB3 tracks and a visual EB3 track overlay (Figure 2C-F). Track overlays are color-coded and color distinctions can be determined by parameters entered by the user. Commonly, these distinctions use the global mean of MT growth speed and MT growth lifetime5,31 and thereby split the data into four groups: slow and short-lived (red), slow and long-lived (yellow), fast and short-lived (green), and fast and long-lived MT growth (blue; Figure 2G). PlusTipTracker-mediated tracking of EB3-labeled MT growth dynamics also enables user-defined Regions of Interest (ROIs). PlusTipTracker extracts the MT growth track data from within the user-defined ROI (Figure 2A, B and F), thereby allowing for comparisons of MT growth dynamics within different regions of the same cell.
<img alt="Figure 2" src="/files/ftp_upload/54265/54265fig2.jpg" /> 
Figure 2: EB3 comet detection, tracking, and plusTipTracker data output. (A) time-lapse movie of a HUVEC expressing mApple EB3. Left panel shows a whole cell view and whole cell mask (white line) demarking the cell boundaries. Red box represents the zoomed region shown in the time-lapse images (right panels). Right panels show raw grey-scale time-lapse images of EB3 comets in successive imaging frames (t = 0 sec - t = 12 sec). (B) plusTipTracker detection of EB3 comets from the images shown in (A). Left panel shows a whole cell view of detected comets (yellow dots). Red box represents the zoomed region shown in the time-lapse images (right panels). Right panels highlight five detected EB3 comets (red arrows) and trace those five comets over the 12 sec imaging period. Green dots represent detected EB3 comets that did not exist (or were not detected) in the previous time interval. (C) Maximum intensity projection of 61 frames (2 min movie) of the EB3 comets shown in (A). (D) plusTipTracker MT track overlay of the same 61 frames (2 min movie) shown in (C). (E) Zoom of maximum intensity projection (red box in C). (F) Zoom of plusTipTracker MT track overlay of maximum intensity projection (red box in D). (G) Color-coded legend for track overlays shown in (D) and (F). Designation as slow, fast, short-lived or long-lived is based on the mean value for all growth tracks measured in a group of ten mApple-EB3 expressing HUVECs. For easier visualization, track overlays in (D) and (F) are generated using inverted pixel intensities of the cell image shown in (A). Scale bars = 20 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
HUVEC branching and migration patterns are sensitive to mechano-physical signals from the ECM.
HUVECs are well known to respond to various types of signaling cues and, in doing so, to undergo morphological alterations that dictate an appropriate cellular response to the particular signaling cue(s)36,40-42. HUVECs cultured on fibronectin coated glass substrates (Figure 3A), or on stiff 2-dimensional polyacrylamide ECMs (55 kPa) typically display a round or elongated morphology with few distinct branched protrusions (Figure 3B). However, when cultured on very soft 2-dimensional polyacrylamide ECMs (0.7 kPa), HUVECs display a highly branched morphology that is known to result from compliance-induced downregulation of myosin-II contractility4,5,31 (Figure 3C). Thus, HUVECs regulate their cellular morphology through ECM mechanosensing, and this is accomplished through localized modulation of MT dynamics and acto-myosin contractility.
Because HUVECs display a wide variety of morphological attributes, in order to measure HUVEC branching it is critical to first define what a branched protrusion is, and then to define how a branched protrusion will be measured objectively. One technique uses a four-step approach by which a cell boundary or &#34;mask&#34; is generated by hand tracing the edges of the cell using an image of a cell expressing a fluorescent volume marker to define the cell edges (Figure 3D). Following mask generation, branch origins are defined by locating greatest angle of curvature between the branched protrusion and the cell body. A line is drawn to demark the &#34;branch origin&#34; (purple angled lines; Figure 3E). Branch number is measured simply by counting the number of branches with a defined branch origin (Figure 3F). Branch length is measured as the distance from the branch origin to the distal branch tip (Figure 3G). To avoid inclusion of small lamellipodial projections from data analysis, additional criterion require that branches must be longer (&#34;branch length&#34;) than they are wide (at the &#34;branch origin&#34;), and that branches must be at least ten microns in length5,31.
<img alt="Figure 3" src="/files/ftp_upload/54265/54265fig3.jpg" /> 
Figure 3: Analysis of HUVEC branching morphology. (A-C) DIC image examples of HUVECs cultured on fibronectin coated glass ECMs (A) stiff (55 kPa) polyacrylamide ECMs (B) and soft (0.7 kPa) polyacrylamide ECMs (C). Branching is dramatically increased in HUVECs cultured on 0.7 kPa polyacrylamide ECMs (C) as compared to stiffer ECMs (A, B). (D). DIC microscopic image (left image) of a transfected HUVEC expressing a mApple-C1 cDNA construct (a cell volume marker; right image). (E) Define the branch origin by locating the greatest angle of curvature (i.e. the position where the membrane displays the greatest curvature; purple-colored lines) on either side of the branch and then connect the angles with a straight line (blue lines). (F) Identify and selecting branched protrusions that extend &#62;10 &#956;m from the &#34;branch origin&#34; (defined in E). Count the number of branched protrusions to obtain the &#34;branch number.&#34; (G) Measure the distance from the center of the &#34;branch origin&#34; to the most distal point of the branch using the line tool in the annotations and measurements application of NIS Elements Software to obtain the &#34;branch length.&#34; Scale bars = 20 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In addition to sensing ECM compliance, HUVECs also sense and respond to the elimination of adjacent cellular contacts as induced by scratch wounding. In the migration assay, HUVECs cultured in a monolayer are subjected to a scratch wound and migration characteristics are measured (see Protocol 9, above). After induction of the wound by scratching the monolayer, wound-edge HUVECs polarize by developing a protrusive leading edge that extends into the wound (Figure 4A, t = 0 hr). Over a time course of 10-12 hr, polarized wound-edge HUVECs migrate into the wound area and fill the wound, establishing a new confluent monolayer (Figure 4A, t = 11 hr). As with HUVEC branching, current evidence suggests that polarization and migration is critically dependent upon the coordinated re-organization of the MT and actin cytoskeletons43-49. This is achieved, in part, through localized inhibition of MCAK-induced MT disassembly at the leading edge, but not at the trailing edge of wound-edge HUVECs31. The sensitivity of MT dynamics to wound-induced polarization is initiated by the Rac1 GTPase, which targets numerous downstream regulator proteins, including Aurora-A kinase that phosphorylates and inhibits MCAK at the leading edge, and other kinases that are capable of inhibiting or activating MAPs to locally modulate MT growth and shrinkage18,31,36,48,50-53. Extensive experimental evidence supports the notion that coordination of leading edge protrusion and trailing edge contraction is governed through cycles of Rac1 activation to promote MT assembly at the leading edge and RhoA activation to acto-myosin contractility at the trailing edge, thereby driving directed motility9,41,51,52,54-58.
To date, there is not an established methodology for performing automated tracking of cell migration. This is a result of difficulty in computationally analyzing cell-ECM boundaries that are consistently changing as wound-edge cells polarize and migrate. While many software programs are capable of tracking fluorescently labeled cells59-64, the population of fluorescently labeled HUVECs in wound-edge migration experiments is approximately 30-40% of the total HUVEC population, and therefore the majority of the wound-edge cells must be visualized by phase-contrast or DIC imaging. Additionally, it is important when measuring wound-edge migration to evaluate fluorescent and non-fluorescent cells within the same wound in order to identify specific effects of expression constructs both within experimental study groups and between experimental study groups. Currently, the best approach to measure HUVEC migration is to use a hand tracking approach in which a wound-edge cell is first identified and then the nucleus and nucleolus are identified (Figure 4B). The nucleolus is used as the position determinant for cell tracking due to its high light scattering density and relatively small size. These two features of the nucleolus make it easier to identify and reduce measurement error by limiting the area in which the user can place individual track positions. HUVEC migration tracks are then generated by clicking on the nucleolus in successive time-lapse image frames. Finally, migration velocity and migration distance to origin are measured and analyzed for control and experimental cell groups (Figure 4B).
<img alt="Figure 4" src="/files/ftp_upload/54265/54265fig4.jpg" /> 
Figure 4: Analysis of wound-edge HUVECs and migration tracking. (A) 20X phase-contrast images of an 11 hr time-lapse imaging series of scratch wound-edge HUVECs (a.k.a. &#34;Migration Assay&#34;). The wound and direction of migration are indicated in panel t = 0 hr. HUVECs are shown to cross the wound-edge and advance into the wound until a monolayer of cells is formed (t = 3.5 hr - t = 11 hr). (B) Analysis of migration assay time lapse imaging (as shown in A) requires first the identification of a trackable wound-edge HUVEC based on cell location (i.e. cells physically at the edge of the wound) and expression profile (i.e., select a green wound edge cell if the experiment involves expression of GFP). Using the phase contrast image, identify the nucleus and the nucleolus. Hand track the nucleolus in successive time-lapse images using the annotations and measurements application in NIS Elements software to determine migration velocity and distance to origin. Scale bars = 100 &#956;m&#160; (A), 20 &#956;m (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells at JoVE.com

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

high-resolution-time-lapse-imaging-automated-analysis-microtubule

Nanyang Technological University

Research

JoVE Journal

Biology

11.0K Views.  University of the Sciences in Philadelphia. Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed. 

Video: High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

Isolation of Human Umbilical Vein Endothelial Cells (HUVEC)

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps include: degradation of the basement membrane, proliferation and migration (sprouting) of endothelial cells (EC) into the extracellular matrix, alignment of EC into cords, lumen formation, anastomosis, and formation of a new basement membrane. Many in vitro assays have been developed to study this process, but most only mimic certain stages of angiogenesis, and morphologically the vessels often do not resemble vessels in vivo. Here we demonstrate an optimized in vitro angiogenesis assay that utilizes human umbilical vein EC and fibroblasts. This model recapitulates all of the key early stages of angiogenesis, and importantly the vessels display patent intercellular lumens surrounded by polarized EC. Vessels can be easily observed by phase-contrast and time-lapse microscopy, and recovered in pure form for downstream applications.

Isolation of Human Umbilical Vein Endothelial Cells HUVEC

This video protocol illustrates the isolation and culture of human umbilical vein endothelial cells (HUVEC) from human umbilical cord. Once isolated these cells can be used for in vitro angiogenesis assays like the Optimized Fibrin Gel Bead Assay also demonstrated by the Hughes lab.

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During directional migration, cells move rapidly in response to an extracellular chemotactic signal, or in response to intrinsic cues 3 provided by the basic motility machinery. Random migration occurs when a cell possesses low intrinsic directionality, allowing the cells to explore their local environment.
Cell migration is a complex process, in the initial response cell undergoes polarization and extends protrusions in the direction of migration 2. Traditional methods to measure migration such as the Boyden chamber migration assay is an easy method to measure chemotaxis in vitro, which allows measuring migration as an end point result. However, this approach neither allows measurement of individual migration parameters, nor does it allow to visualization of morphological changes that cell undergoes during migration.
Here, we present a method that allows us to monitor migrating cells in real time using video - time lapse microscopy. Since cell migration and invasion are hallmarks of cancer, this method will be applicable in studying cancer cell migration and invasion in vitro. Random migration of platelets has been considered as one of the parameters of platelet function 4, hence this method could also be helpful in studying platelet functions. This assay has the advantage of being rapid, reliable, reproducible, and does not require optimization of cell numbers. In order to maintain physiologically suitable conditions for cells, the microscope is equipped with CO2 supply and temperature thermostat. Cell movement is monitored by taking pictures using a camera fitted to the microscope at regular intervals. Cell migration can be calculated by measuring average speed and average displacement, which is calculated by Slidebook software.

This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During ...

Phenotypic and Functional Characterization of Endothelial Colony Forming Cells Derived from Human Umbilical Cord Blood

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of circulating endothelial progenitor cells (EPCs) were first identified in adult human peripheral blood by Asahara et al. in 1997 2 bringing an infusion of new hypotheses and strategies for vascular regeneration and repair. EPCs are rare but normal components of circulating blood that home to sites of blood vessel formation or vascular remodeling, and facilitate either postnatal vasculogenesis, angiogenesis, or arteriogenesis largely via paracrine stimulation of existing vessel wall derived cells3. No specific marker to identify an EPC has been identified, and at present the state of the field is to understand that numerous cell types including proangiogenic hematopoietic stem and progenitor cells, circulating angiogenic cells, Tie2+ monocytes, myeloid progenitor cells, tumor associated macrophages, and M2 activated macrophages participate in stimulating the angiogenic process in a variety of preclinical animal model systems and in human subjects in numerous disease states4, 5. Endothelial colony forming cells (ECFCs) are rare circulating viable endothelial cells characterized by robust clonal proliferative potential, secondary and tertiary colony forming ability upon replating, and ability to form intrinsic in vivo vessels upon transplantation into immunodeficient mice6-8. While ECFCs have been successfully isolated from the peripheral blood of healthy adult subjects, umbilical cord blood (CB) of healthy newborn infants, and vessel wall of numerous human arterial and venous vessels 6-9, CB possesses the highest frequency of ECFCs7 that display the most robust clonal proliferative potential and form durable and functional blood vessels in vivo8, 10-13. While the derivation of ECFC from adult peripheral blood has been presented14, 15, here we describe the methodologies for the derivation, cloning, expansion, and in vitro as well as in vivo characterization of ECFCs from the human umbilical CB.

Endothelial colony forming cells (ECFCs) are circulating endothelial cells with robust clonal proliferative potential that display intrinsic in vivo vessel forming ability. Phenotypic and functional characterization of outgrowth endothelial cells derived from CB are important to identify and isolate bona fide ECFCs for potential clinical application in repairing damaged tissues.

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.

We present a simple protocol to obtain fluorescence microscopy movies of growing yeast cells, and a GUI-based software package to extract single-cell time series data. The analysis includes automated lineage and division time assignment integrated with visual inspection and manual curation of tracked data.

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies ...

Analysis of Oxidative Stress in Zebrafish Embryos

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes oxidation of molecules (lipid, DNA, protein) and leads to cell death. Oxidative stress also impacts the progression of several pathological conditions such as diabetes, retinopathies, neurodegeneration, and cancer. Thus, it is important to define tools to investigate oxidative stress conditions not only at the level of single cells but also in the context of whole organisms. Here, we consider the zebrafish embryo as a useful in vivo system to perform such studies and present a protocol to measure in vivo oxidative stress. Taking advantage of fluorescent ROS probes and zebrafish transgenic fluorescent lines, we develop two different methods to measure oxidative stress in vivo: i) a &#8220;whole embryo ROS-detection method&#8221; for qualitative measurement of oxidative stress and ii) a &#8220;single-cell&#160;ROS detection method&#8221; for quantitative measurements of oxidative stress. Herein, we demonstrate the efficacy of these procedures by increasing oxidative stress in tissues by oxidant agents and physiological or genetic methods. This protocol is amenable for forward genetic screens and it will help address cause-effect relationships of ROS in animal models of oxidative stress-related pathologies such as neurological disorders and cancer.

Here we report a protocol to measure oxidative stress in living zebrafish embryos. This procedure allows reactive oxygen species (ROS) detection in both whole embryo tissues and single-cell populations. This protocol will accomplish both qualitative and quantitative analyses.

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and several essential mitochondrial polypeptides. Mitochondrial nucleoids divide and distribute within the dynamic mitochondrial network that undergoes fission/fusion and other morphological changes. High resolution live fluorescence microscopy is a straightforward technique to characterize a nucleoid&#39;s position and motion. For this technique, nucleoids are commonly labeled through fluorescent tags of their protein components, namely transcription factor a (TFAM). However, this strategy needs overexpression of a fluorescent protein-tagged construct, which may cause artifacts (reported for TFAM), and is not feasible in many cases. Organic DNA-binding dyes do not have these disadvantages. However, they always show staining of both nuclear and mitochondrial DNAs, thus lacking specificity to mitochondrial nucleoids. By taking into account the physico-chemical properties of such dyes, we selected a nucleic acid gel stain (SYBR Gold) and achieved preferential labeling of mitochondrial nucleoids in live cells. Properties of the dye, particularly its high brightness upon binding to DNA, permit subsequent quantification of mitochondrial nucleoid motion using time series of super-resolution structured illumination images.

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and ...