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Real‐Time Centrosome Reorientation During Fibroblast Migration
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[45] Real‐Time Centrosome Reorientation During Fibroblast Migration By EDGAR R. GOMES and GREGG G. GUNDERSEN Abstract
The centrosome is positioned between the nucleus and the leading edge of many types of migrating cells. Cdc42 regulates this centrosome reorientation through its effectors Par6 and MRCK. Using time‐lapse microscopy of live cells, the mechanisms and kinetics of centrosome reorientation can be studied. In this chapter, we describe a modification in the standard wound healing assay that allows the study of signaling pathways involved in centrosome reorientation and other polarization events that occur before cell migration. We also describe a method for visualization of centrosome reorientation by time‐lapse microscopy using NIH 3T3 fibroblasts stably transfected with GFP‐tubulin. Introduction
During migration of many cell types, the centrosome is reoriented to a position between the nucleus and the leading edge. The centrosome is the microtubule (MT) nucleating site, and in most migrating cells, the MTs remain attached to the centrosome, giving rise to a radial array of MTs. Because of the organizing capability of the centrosome, it is sometimes referred to as the MT organizing center (MTOC), and centrosome reorientation is also referred to as MTOC reorientation. Centrosome reorientation toward the leading edge has been described in migrating fibroblasts, neurons, astrocytes, endothelial cells, and macrophages (Etienne‐Manneville and Hall, 2001; Euteneuer and Schliwa, 1992; Gotlieb et al., 1981; Gregory et al., 1988; Gundersen and Bulinski, 1988; Kupfer et al., 1982; Nemere et al., 1985). Centrosome reorientation occurs in some nonmigratory situations such as in endothelial cells in situ (Rogers et al., 1985) or in response to shear stress in culture (Tzima et al., 2003) and in T cells, where the centrosome is reoriented toward the target cell (Kupfer et al., 1983). The reorientation of the centrosome and the associated Golgi apparatus may facilitate the delivery of components to the leading edge (Bergmann et al., 1983; Prigozhina and Waterman‐Storer, 2004; Schmoranzer et al., 2003). Centrosome reorientation may also contribute to the appropriate geometry to ensure that the nucleus moves forward with
METHODS IN ENZYMOLOGY, VOL. 406 Copyright 2006, Elsevier Inc. All rights reserved.
0076-6879/06 $35.00 DOI: 10.1016/S0076-6879(06)06045-9
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the cell (Gomes et al., 2005; Solecki et al., 2004). The molecular mechanisms of centrosome reorientation are not known, however, in several systems, Cdc42 and its effectors, Par6 and MRCK, and dynein have been implicated in the regulation of centrosome reorientation (Etienne‐ Manneville, 2004; Gomes et al., 2005). Centrosome reorientation in migrating cells is easily quantified in fixed preparations stained with centrosomal and nuclear markers. A cell is determined to have a reoriented centrosome when the centrosome is located in the ‘‘pie slice’’ defined by the nucleus and the leading edge (Fig. 1). As this sector occupies 1/3 of the cell, by chance, 33% of the
FIG. 1. (A) Diagram of criteria used to determined centrosome orientation. When the centrosome is localized in the sector between the nucleus and the leading edge, it is scored as oriented. Localization in other sectors is scored as non‐oriented. (B, C) Wound edge cells with oriented and non‐oriented centrosomes. Cells were treated with LPA, fixed, and stained for MTs (B) and pericentrin, a centrosomal protein (C). In both images, the nucleus is detected as an oval nonfluorescent area. Cells with (þ) and without () reoriented centrosomes are indicated in C.
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cells will have a ‘‘reoriented centrosome.’’ In wounded monolayers, which are typically used to study centrosome reorientation, maximal centrosome reorientation reaches 70–90% and occurs 2 h after wounding of endothelial and fibroblast monolayers (Gundersen and Bulinski, 1988) and 8 h after wounding astrocyte monolayers (Etienne‐Manneville and Hall, 2001). Determination of centrosome reorientation in fixed‐cell preparations is a simple assay. However, it does not provide information on the mechanism, time, and speed of centrosome reorientation or what actually moves during centrosome reorientation. A number of techniques have been used to visualize centrosomes in living cells so that these issues can be addressed. Stable cell lines expressing green fluorescent protein (GFP)‐tagged ‐tubulin, a centrosomal protein, have characterized centrosome position during migration of CHO and Ptk cells (Yvon et al., 2002). A transgenic mouse expressing GFP‐centrin‐2 has been described, and it will be a valuable tool to studycentrosome reorientation by time‐lapse microscopy (Higginbotham et al., 2004). Centrosome position can also be determined by visualizing the focus of MTs (i.e., by the position of the MTOC) (Gomes et al., 2005). MTs visualized by modulated polarization microscopy have been used to visualize centrosome reorientation in T cells (Kuhn and Poenie, 2002). In this chapter, we describe the preparation and use of stably expressing GFP‐‐tubulin NIH 3T3 cell line (3T3‐GFPTub) to observe centrosome reorientation in wounded monolayers (Gomes et al., 2005). As the GFP‐ tubulin is incorporated into both centriolar and cytoplasmic MTs, the dynamics of the centrosome and cytoplasmic MTs can be followed simultaneously. This allows movements of the centrosome to be related to changes in MT dynamics and/or interactions with cortical sites. Another advantage of using such a cell line is that the position of the nucleus can be determined, because the nucleus is easily identified in the GFP‐tubulin–expressing cell as a nonfluorescent area. In our experience, transient expression of GFP‐ tubulin has a number of drawbacks: there is substantial cell–cell variability in expression levels, and high levels of GFP‐tagged tubulin can alter MT dynamics and give rise to substantial diffuse cytoplasmic fluorescence. MTs can also be labeled by microinjection of fluorescently labeled tubulin (Waterman‐Storer, 2002). This method requires a microinjection apparatus and can be more time consuming and technically demanding. Generation of a Stable Cell Line Expressing GFP‐Tubulin
We used a standard method for generating stable NIH‐3T3 cell lines expressing GFP‐tubulin (Sullivan and Satchwell, 2000). Cells are transfected with a commercially available enhanced GFP‐‐tubulin vector containing a neomycin‐resistance cassette (BD Biosciences Clontech).
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Addition of neomycin or G418 (a neomycin analog) selects for cells that incorporate the vector. Resistant clones are screened for GFP‐‐tubulin expression levels and checked for growth rates, migration, centrosome reorientation, and formation of stable MTs (Cook et al., 1998; Gomes et al., 2005). Materials NIH 3T3 cells (ATCC) DMEM (Invitrogen) Enhanced GFP‐‐tubulin mammalian expressing vector (Clontech) Lipofectamine (Invitrogen) G418 (Calbiochem) Cloning cylinders (Sigma) NIH 3T3 fibroblasts are cultured in DMEM supplemented with 10% calf serum (Cook et al., 1998; Gundersen et al., 1994) and kept at subconfluent densities to prevent spontaneous transformation. Transfect cells with GFP‐‐tubulin vector using lipofectamine according to the protocol provided by the manufacture. Two days after transfection, plate cells at low density with DMEM medium supplemented with 1 mg/ml G418. Once colonies are visible by direct observation of the plates (5–7 days after transfection), select individual colonies using cloning cylinders and transfer them to larger plates for expansion. Reduce G418 concentration to 0.5 mg/ml at this stage. When clones have been expanded, check for GFP‐‐tubulin expression by plating cells on acid‐washed glass coverslips, fixing in 20 methanol, rehydrating, and observing by fluorescence microscopy. Select clones with fluorescent MTs that are in a normal radial arrays (Fig. 1B,C). Characterize positive clones by growth rate, migration into the wound, and centrosome reorientation (Fig. 2). We also check clones for formation of stabilized MTs, because this is another rearrangement of MTs that occurs in cells in response to wounding (Cook et al., 1998; Gundersen and Bulinski, 1988). Select clones that behave similarly to parental cells. The level of GFP‐‐tubulin relative to endogenous ‐tubulin is <5% in the selected clone (3T3‐GFPTub) (Fig. 2). Multiple clones should be checked for use in cell assays to ensure that the behavior being studied is typical.
Wound Healing Assay
Wound healing assays are useful to study cell polarization and migration. When a confluent monolayer of cells is wounded, the cells at the wound edge synchronously polarize and begin migrating in the direction
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FIG. 2. Characterization of the NIH 3T3 cell line stably expressing GFP‐‐tubulin (3T3 GFPTub). (A) GFP‐tubulin fluorescence in 3T3‐GFPTub cells. (B) Higher magnification of the inset in (A). The different focal plane shows the MTs of the mitotic spindle. Centrosome reorientation stimulated by LPA or serum is identical in parental NIH 3T3 (C) and 3T3 GFPTub (D) cells. Serum‐starved cells were left in serum‐free DMEM (SF) or treated with LPA or calf serum for 2 h. Cells were fixed with methanol, immunofluorescently stained for centrosomes and MTs, and the extent of centrosome reorientation determined. (E) Expression levels of GFP‐‐tubulin in 3T3‐GFPTub cells determined by Western blot using an anti‐‐tubulin antibody. GFP‐‐tubulin is expressed at <5% the level of endogenous ‐tubulin.
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of the wound. For cells maintained in serum, the polarization and migration responses are triggered as a result of wounding. We introduced a small variation in the wounding assay that allows us to trigger wound response with soluble factors rather than by wounding alone. If confluent monolayers of NIH 3T3 cells are serum starved before wounding, the cells at the edge of the wound do not polarize or migrate until the appropriate serum factors are added (Fig. 3) (Gundersen et al., 1994). The mitogenic lipid lysophosphatidic acid (LPA) is a major serum factor required for MT polarizations in response to wounding (Cook et al., 1998; Palazzo et al., 2001). LPA induces the same MT reorganizations as serum, yet LPA alone is not sufficient to stimulate cell migration (Gomes et al., 2005). Thus, addition of LPA only activates cell polarization and allows the study of polarization events independently from those of active cell migration. To induce both cell polarization and migration responses, serum can be added to the starved cells (Gomes et al., 2005; Gundersen and Bulinski, 1988; Gundersen et al., 1994). Because serum factors are necessary for triggering the responses to wounding, serum starvation also allows a separation of the physical act of wounding from the triggering of polarization and migration responses. This has a number of advantages for studying cell polarization and migration, particularly in studying the very early polarization and migration responses. For example, purified proteins, antibodies, or expression plasmids can be introduced into wound edge cells by microinjection before stimulation, which is not possible with the regular wounding assay.
FIG. 3. Wound assay using serum‐starved monolayers of NIH 3T3 fibroblasts. Schematic representation of a serum‐starved monolayer with MTs, centrosome and nucleus. When the starved monolayer is wounded, centrosome reorientation does not occur. Centrosome reorientation is triggered only on the addition of LPA or serum. (See Palazzo et al. [2001] for details.)
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Introduced proteins can also be tested for sufficiency in induction of cell polarization, which is not possible in a regular wounding assay (Gomes et al., 2005; Palazzo et al., 2001; Wen et al., 2004). We have also found that serum‐starved cells are flatter and provide better subjects for optical imaging. In the following, we describe the specific protocols for serum starvation and wounding of NIH 3T3 fibroblasts. We have found that similar protocols work well for other rodent fibroblast cell lines (Swiss 3T3, Balb 3T3, and NRK fibroblasts). Materials Glass coverslips, washed with 1 N HCl Jeweler’s screwdriver DMEM medium NIH‐3T3 cells Viokase or trypsin/EDTA LPA (1 mM) Preparation of LPA Stock Solution. Add 2.3 ml of a sterile 10 mM HEPES, pH 7.4, 10 mg/ml BSA (fatty acid free, Sigma), 100 mM NaCl solution to 1 mg of LPA (Sigma). Mix well. Aliquot and freeze at 80 . Frozen stocks are good for at least 2 months. Thawed aliquots remain usable for 24 h. Preparation of Starved Cell Monolayers and Wounding. Cover the bottom of a 10‐cm tissue culture dish with coverslips. Detach NIH 3T3 cells from a stock plate using trypsin/EDTA or Viokase and resuspend in DMEM with 10 % calf serum. Dilute cells into 10 ml of DMEM plus 10% calf serum and add 10 ml per 10‐cm dish so that cells will be approximately 20% confluent. Allow cells to grow until just confluent (48 h). Fill three 6‐cm dishes with 8 ml of DMEM without serum. Fill 35‐mm dishes with 1 ml of DMEM without serum (or each well of a 6‐well plate). Using a sterile forceps, transfer each coverslip (containing the confluent monolayer of cells) serially to the 6‐cm dishes containing DMEM. Place the coverslip in the 35‐mm dish and starve the cells for 48 h. Some cells will detach during starvation; however, the remaining cells should still form a confluent monolayer and are fully viable (Gundersen et al., 1994). Using a sterile jeweler’s screwdriver (tip width 1–2 mm), wound the monolayer with an even, continuous movement. Multiple wounds can be made on a single coverslip. Replace cells in incubator and let recover for 20 min. Treat cells with 2 M LPA to induce centrosome reorientation and MT stabilization or add serum (1–5%) to stimulate migration. Centrosome reorientation, MT stabilization, and cell migration can be assessed with fixed cell
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assays (see Cook et al. [1998]; Palazzo et al. [2001]; Wen et al. [2004]), or 3T3‐GFPTub cells can be used for live cell imaging (see the following). Note Wound edge cells can be microinjected after recovery from the wound. Proteins can be microinjected into the cytoplasm, and DNA can be injected into the nucleus. Typically, we find that DNA expression vectors based on CMV promoters allows for efficient protein expression in 1–2 h after microinjection. Because cells will still polarize and migrate if LPA or serum are added up to 8 h after wounding, this allows proteins to be expressed before triggering wound responses. Imaging Centrosome Reorientation in Living Cells
For real‐time imaging of centrosome reorientation, we use 3T3‐ GFPTub cells prepared as described previously. The preparation of the 3T3‐GFPTub cell monolayers for wounding and serum starvation is similar to the preparation of NIH 3T3 cells for wound healing assays (see preceding) with a few modifications to allow for optimal imaging and maintenance of cells under the microscope. 3T3‐GFPTub cells are plated in specific imaging chambers, and modified medium (‘‘recording medium’’) is used for starvation and for maintaining the cells while imaging. Materials Inverted microscope equipped with epi‐fluorescence (Nikon Eclipse TE300) 60 Plan Apo (NA 1.4) objective 40 Plan Fluor (NA 0.6) phase objective Computer‐controlled shutters (Sutter and Uniblitz) on the epi‐illumination light path and the bright‐field illumination light path (for phase‐ contrast imaging) Neutral density filters (ND4 and ND8) on the epi‐illumination light path GFP‐optimized filter set (Chroma) Temperature‐controlled chamber surrounding the microscope Motorized Z‐focus (Nikon) Motorized XY stage (Prior) Cooled CCD camera (Coolsnap HQ)
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Computer with software to control shutters, time‐lapse acquisition, Z‐focus, and XY stage (Metamorph, Universal Imaging Corporation) Recording medium Glass coverslip chambers Preparation of Recording Medium. HBSS (Gibco) containing essential and nonessential MEM amino acids (Gibco), 2.5 g/l D‐glucose, 2 mM glutamine, 1 mM sodium pyruvate, and 10 mM HEPES, pH 7.4. Preparation of Glass Coverslip Imaging Chambers. 35‐mm plastic tissue culture dishes are used. A hole (10–15 mm diameter) is punched in the bottom of the dish with a hand punch tool (Roper Whitney). An acid‐ washed glass coverslip (22 mm2 ) is glued to the bottom of the dish, covering the hole, using Sylgard 184 Silicone Elastomer (Dow Corning) or VALAP (Vaseline/lanolin/paraffin ¼ 1:1:1). Plates are sterilized under UV light overnight. We have also had good results with commercially prepared glass coverslip dishes (35‐mm dish with 10‐mm hole size) from Mattek. Live Cell Imaging of Centrosome Reorientation. Plate 3T3‐GFPTub cells in imaging chambers at a 20% density (see preceding). Once cells reach confluency (after 48 h), wash the cells twice with recording medium and add 4 ml of recording medium. Starve cells in recording medium for 24 h. Equilibrate the microscope temperature chamber at 34–35 for at least 1 h before recording. Wound the monolayer and allow to recover for 20 min in the incubator. Transfer the imaging chamber to the microscope stage. To minimize photodamage (see following), insert the highest neutral density filter in the epi‐illumination light path that still allows observation of the centrosome. Select cells with distinct centrosome and MTs. Minimize as much as possible the exposure of cells to epi‐fluorescence illumination. Determine the minimum exposure time necessary for obtaining an image with a distinct centrosome (see following). Start image acquisition at a rate that does not induce photobleaching (see following) to characterize centrosome position before stimulation. To induce centrosome reorientation, add 100 l of 80 M LPA drop‐wise (2 M final concentration) or 40 l of serum (1% final concentration). Restart image acquisition. We have successfully followed centrosome position up to 4 h after addition of LPA, acquiring 1 image per minute, or up to 18 h after addition of serum, acquiring 1 image every 10 min. By use of this protocol, we were able to observe that centrosome reorientation occurs by nuclear movement away from the leading edge, whereas the centrosome remains at the cell centroid (Fig. 4). If additional information about the nucleus or the cell boundaries are required (for example, to relate centrosome reorientation to leading edge
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FIG. 4. High‐magnification imaging of centrosome and nucleus position during LPA‐ stimulated centrosome reorientation. Selected frames from a time‐lapse movie of a serum‐ starved 3T3‐GFPTub cell before (A) and after (C) the addition of LPA at 0:26 (h:min). The centrosome is identified as a bright point at the center of the MT array. The wound is at the top of the panel. The nucleus is the nonfluorescent oval area. For each time‐point, the position of the centrosome and the centroid of the nucleus were determined and plotted before (B) and after (D) the addition of LPA. Before the addition of LPA, both centrosome and LPA remain in the same position. After the addition of LPA, the nucleus moves away from the leading edge, whereas the centrosome remains stationary at the cell center. Thus, the rearward movement of the nucleus reorients the centrosome. Images were acquired with a 60 Plan Apo (NA 1.4) objective, ND8 density filter, camera binning of 2 2, and 1500 msec exposure at a rate of 1 frame/min. Previously published in Cell (Gomes et al., 2005).
structures, such as lamellipodia or ruffles) sequential phase and fluorescent images can be acquired. To do this, the microscope needs to be equipped with phase or differential interference contrast (DIC) objectives and a shutter in the transmitted light path. The phase or DIC image also contains information regarding possible rotation of the nucleus by reference to the position of the nucleolus and other structures within the nucleus. Metamorph and other commercially available software can be programmed to acquire both epifluorescence and phase images at each time point.
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Notes Chamber Temperature. Preheating the microscope incubator and increasing room temperature minimizes focal drift. We use a temperature of 34–35 instead of 37 , because it is easier to maintain this temperature and focal drift is less of a problem. Recording Media. DMEM medium is not suitable for fluorescence microscopy because it contains riboflavin that interferes with GFP fluorescence microscopy. Riboflavin has an excitation peak at 450 nm and an emission peak at 530 nm, which is similar to enhanced GFP excitation and emission peaks (480 nm and 530 nm, respectively). We also leave out the phenol red that is commonly included in media, because it can absorb light, and this is important for fluorochromes such as rhodamine or DsRed. The recording medium is buffered with HEPES, not bicarbonate, because most microscope chambers are not equipped with a CO2‐enriched atmosphere. The imaging chamber must be covered, because evaporation is significant at 37 , especially during long recordings. Starvation. Starvation time is reduced to 24 h when recording medium is used, because cells start to detach from the coverslip if they are starved for longer times in recording medium. Alternately, cells can be starved with DMEM for 48 h, wounded, transferred to the microscope, and then stimulated with recording medium containing LPA or serum. Starved cells are sensitive to changes in medium; therefore, if the starvation medium is replaced by recording medium without LPA or serum, cells can detaching rapidly (15–30 min). Imaging of the centrosome position before stimulation is not possible if cells are starved in DMEM. Photodamage. Long exposure times can result in photodamage of the cells without detectable photobleaching of the GFP‐tubulin. Photodamage is recognized when cells bleb, MTs depolymerize or, more subtly, do not respond to LPA or serum addition. To minimize photodamage, reduce light exposure as much as possible while searching for cells and during imaging. It is helpful to dark‐adapt one’s eyes by waiting 5 min in complete darkness before searching for cells under the microscope. Also, we have found that the use of neutral density filters in combination with longer exposure times generates less photodamage than short exposure times without neutral density filters. Image Acquisition. Imaging centrosome reorientation can involve long (up to 4 h) recording times. To prevent photobleaching and photodamage, we acquired images at a rate between 1 frame per 1 min and 1 frame every 10 min. These acquisition rates are sufficient to characterize both centrosome and nucleus movements. The exposure used for
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image acquisition should be as short as possible without compromising the visualization of the centrosome. Setting the camera binning to 2 2, acquisition time can be shortened; however, image resolution is also reduced. Triggering Centrosome Reorientation. Centrosome reorientation can be triggered either by LPA or serum, as described previously. The concentration of serum should not be higher that 1%, because serum contains fluorescent factors that increase the background fluorescence, decreasing image quality. This amount of serum is sufficient to induce cell polarization and subsequent migration (Fig. 5). Objectives. We find that a 60 oil Plan Apo objective (NA 1.4) gives the best combination of resolution, sensitivity, and field size for imaging centrosome reorientation. With this objective, the centrosome appears distinct, and individual MTs can be observed (Fig. 4). With a higher magnification objective (e.g., 100), only part of the cell is observed, and photobleaching occurs faster. When only the position of the centrosome
FIG. 5. Low‐magnification imaging of centrosome and nucleus position during serum‐ stimulated centrosome reorientation. Selected frames from a time‐lapse movie of a starved 3T3‐GFPTub cell after the addition of calf serum at 0:00 (min:sec). In this example, a 40 phase objective was used, and both phase‐contrast (top) and epifluorescence (bottom) images were acquired. Although it is not possible to observe individual microtubules in the fluorescence image, the centrosome is visible. The position of the centrosome is represented by a white triangle in the phase‐contrast image. Images were acquired with a 40 Plan Fluor (NA 0.6) phase objective, ND8 density filter, camera binning of 1 1, and 500 msec exposure time at a rate of 1 frame every 3 min.
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is to be determined, we obtained good results using a 40 Plan Fluor objective NA 0.6 (Fig. 5). One advantage of using a lower magnification objective is that higher numbers of cells can be imaged. XY Motorized Stage. Using a motorized XY stage, we have been able to acquire multiple fields (up to 60) at each time point (one frame every 3 min), increasing the speed of data acquisition. A motorized Z‐focus and an autofocus controller are necessary for the acquisition of in‐focus images. Autofocusing can be software controlled. The autofocus software routine must be performed using phase‐contrast illumination to prevent photobleaching of GFP; therefore, a shutter in the transmitted illumination path and phase condenser and objectives are required. We find that 40 dry objectives perform optimally in this scenario. Oil immersion objectives can also be used as long as the travel distances between each field are minimized (<1 cm).
Acknowledgments We thank Jan Schmoranzer for comments on the manuscript. This work was supported by a postdoctoral fellowship from Fundac¸ a˜ o para a Cieˆ ncia e Tecnologia, Portugal (to E. R. G.), and an NIH grant GM062938 (to G. G. G.).
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Gundersen, G. G., Kim, I., and Chapin, C. J. (1994). Induction of stable microtubules in 3T3 fibroblasts by TGF‐beta and serum. J. Cell Sci. 107, 645–659. Higginbotham, H., Bielas, S., Tanaka, T., and Gleeson, J. G. (2004). Transgenic mouse line with green‐fluorescent protein‐labeled Centrin 2 allows visualization of the centrosome in living cells. Transgenic Res. 13, 155–164. Kuhn, J. R., and Poenie, M. (2002). Dynamic polarization of the microtubule cytoskeleton during CTL‐mediated killing. Immunity 16, 111–121. Kupfer, A., Louvard, D., and Singer, S. J. (1982). Polarization of the Golgi apparatus and the microtubule‐organizing center in cultured fibroblasts at the edge of an experimental wound. Proc. Natl. Acad. Sci. USA 79, 2603–2607. Kupfer, A., Dennert, G., and Singer, S. J. (1983). Polarization of the Golgi apparatus and the microtubule‐organizing center within cloned natural killer cells bound to their targets. Proc. Natl. Acad. Sci. USA 80, 7224–7228. Nemere, I., Kupfer, A., and Singer, S. J. (1985). Reorientation of the Golgi apparatus and the microtubule‐organizing center inside macrophages subjected to a chemotactic gradient. Cell Motil. 5, 17–29. Palazzo, A. F., Joseph, H. L., Chen, Y. J., Dujardin, D. L., Alberts, A. S., Pfister, K. K., Vallee, R. B., and Gundersen, G. G. (2001). Cdc42, dynein, and dynactin regulate MTOC reorientation independent of Rho‐regulated microtubule stabilization. Curr. Biol. 11, 1536–1541. Prigozhina, N. L., and Waterman‐Storer, C. M. (2004). Protein kinase D‐mediated anterograde membrane trafficking is required for fibroblast motility. Curr. Biol. 14, 88–98. Rogers, K. A., McKee, N. H., and Kalnins, V. I. (1985). Preferential orientation of centrioles toward the heart in endothelial cells of major blood vessels is reestablished after reversal of a segment. Proc. Natl. Acad. Sci. USA 82, 3272–3276. Schmoranzer, J., Kreitzer, G., and Simon, S. M. (2003). Migrating fibroblasts perform polarized, microtubule‐dependent exocytosis towards the leading edge. J. Cell Sci. 116, 4513–4519. Solecki, D. J., Model, L., Gaetz, J., Kapoor, T. M., and Hatten, M. E. (2004). Par6 signaling controls glial‐guided neuronal migration. Nat. Neurosci. 7, 1195–1203. Sullivan, J., and Satchwell, M. F. (2000). Development of stable cell lines expressing high levels of point mutants of human opsin for biochemical and biophysical studies. Methods Enzymol. 315, 30–58. Tzima, E., Kiosses, W. B., del Pozo, M. A., and Schwartz, M. A. (2003). Localized cdc42 activation, detected using a novel assay, mediates microtubule organizing center positioning in endothelial cells in response to fluid shear stress. J. Biol. Chem. 278, 31020–31023. Waterman‐Storer, C. M. (2002). Fluorescent speckle microscopy (FSM) of microtubules and actin in living cell. In ‘‘Current Protocols in Cell Biology’’ (K. S. Morgan, ed.), pp. 4.10.11–14.10.26. John Wiley & Sons Inc., New York. Wen, Y., Eng, C. H., Schmoranzer, J., Cabrera‐Poch, N., Morris, E. J., Chen, M., Wallar, B. J., Alberts, A. S., and Gundersen, G. G. (2004). EB1 and APC bind to mDia to stabilize microtubules downstream of Rho and promote cell migration. Nat. Cell Biol. 6, 820–830. Yvon, A. M., Walker, J. W., Danowski, B., Fagerstrom, C., Khodjakov, A., and Wadsworth, P. (2002). Centrosome reorientation in wound‐edge cells is cell type specific. Mol. Biol. Cell 13, 1871–1880.
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[45] Real‐Time Centrosome Reorientation During Fibroblast Migration By EDGAR R. GOMES and GREGG G. GUNDERSEN Abstract
The centrosome is positioned between the nucleus and the leading edge of many types of migrating cells. Cdc42 regulates this centrosome reorientation through its effectors Par6 and MRCK. Using time‐lapse microscopy of live cells, the mechanisms and kinetics of centrosome reorientation can be studied. In this chapter, we describe a modification in the standard wound healing assay that allows the study of signaling pathways involved in centrosome reorientation and other polarization events that occur before cell migration. We also describe a method for visualization of centrosome reorientation by time‐lapse microscopy using NIH 3T3 fibroblasts stably transfected with GFP‐tubulin. Introduction
During migration of many cell types, the centrosome is reoriented to a position between the nucleus and the leading edge. The centrosome is the microtubule (MT) nucleating site, and in most migrating cells, the MTs remain attached to the centrosome, giving rise to a radial array of MTs. Because of the organizing capability of the centrosome, it is sometimes referred to as the MT organizing center (MTOC), and centrosome reorientation is also referred to as MTOC reorientation. Centrosome reorientation toward the leading edge has been described in migrating fibroblasts, neurons, astrocytes, endothelial cells, and macrophages (Etienne‐Manneville and Hall, 2001; Euteneuer and Schliwa, 1992; Gotlieb et al., 1981; Gregory et al., 1988; Gundersen and Bulinski, 1988; Kupfer et al., 1982; Nemere et al., 1985). Centrosome reorientation occurs in some nonmigratory situations such as in endothelial cells in situ (Rogers et al., 1985) or in response to shear stress in culture (Tzima et al., 2003) and in T cells, where the centrosome is reoriented toward the target cell (Kupfer et al., 1983). The reorientation of the centrosome and the associated Golgi apparatus may facilitate the delivery of components to the leading edge (Bergmann et al., 1983; Prigozhina and Waterman‐Storer, 2004; Schmoranzer et al., 2003). Centrosome reorientation may also contribute to the appropriate geometry to ensure that the nucleus moves forward with
METHODS IN ENZYMOLOGY, VOL. 406 Copyright 2006, Elsevier Inc. All rights reserved.
0076-6879/06 $35.00 DOI: 10.1016/S0076-6879(06)06045-9
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the cell (Gomes et al., 2005; Solecki et al., 2004). The molecular mechanisms of centrosome reorientation are not known, however, in several systems, Cdc42 and its effectors, Par6 and MRCK, and dynein have been implicated in the regulation of centrosome reorientation (Etienne‐ Manneville, 2004; Gomes et al., 2005). Centrosome reorientation in migrating cells is easily quantified in fixed preparations stained with centrosomal and nuclear markers. A cell is determined to have a reoriented centrosome when the centrosome is located in the ‘‘pie slice’’ defined by the nucleus and the leading edge (Fig. 1). As this sector occupies 1/3 of the cell, by chance, 33% of the
FIG. 1. (A) Diagram of criteria used to determined centrosome orientation. When the centrosome is localized in the sector between the nucleus and the leading edge, it is scored as oriented. Localization in other sectors is scored as non‐oriented. (B, C) Wound edge cells with oriented and non‐oriented centrosomes. Cells were treated with LPA, fixed, and stained for MTs (B) and pericentrin, a centrosomal protein (C). In both images, the nucleus is detected as an oval nonfluorescent area. Cells with (þ) and without () reoriented centrosomes are indicated in C.
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cells will have a ‘‘reoriented centrosome.’’ In wounded monolayers, which are typically used to study centrosome reorientation, maximal centrosome reorientation reaches 70–90% and occurs 2 h after wounding of endothelial and fibroblast monolayers (Gundersen and Bulinski, 1988) and 8 h after wounding astrocyte monolayers (Etienne‐Manneville and Hall, 2001). Determination of centrosome reorientation in fixed‐cell preparations is a simple assay. However, it does not provide information on the mechanism, time, and speed of centrosome reorientation or what actually moves during centrosome reorientation. A number of techniques have been used to visualize centrosomes in living cells so that these issues can be addressed. Stable cell lines expressing green fluorescent protein (GFP)‐tagged ‐tubulin, a centrosomal protein, have characterized centrosome position during migration of CHO and Ptk cells (Yvon et al., 2002). A transgenic mouse expressing GFP‐centrin‐2 has been described, and it will be a valuable tool to studycentrosome reorientation by time‐lapse microscopy (Higginbotham et al., 2004). Centrosome position can also be determined by visualizing the focus of MTs (i.e., by the position of the MTOC) (Gomes et al., 2005). MTs visualized by modulated polarization microscopy have been used to visualize centrosome reorientation in T cells (Kuhn and Poenie, 2002). In this chapter, we describe the preparation and use of stably expressing GFP‐‐tubulin NIH 3T3 cell line (3T3‐GFPTub) to observe centrosome reorientation in wounded monolayers (Gomes et al., 2005). As the GFP‐ tubulin is incorporated into both centriolar and cytoplasmic MTs, the dynamics of the centrosome and cytoplasmic MTs can be followed simultaneously. This allows movements of the centrosome to be related to changes in MT dynamics and/or interactions with cortical sites. Another advantage of using such a cell line is that the position of the nucleus can be determined, because the nucleus is easily identified in the GFP‐tubulin–expressing cell as a nonfluorescent area. In our experience, transient expression of GFP‐ tubulin has a number of drawbacks: there is substantial cell–cell variability in expression levels, and high levels of GFP‐tagged tubulin can alter MT dynamics and give rise to substantial diffuse cytoplasmic fluorescence. MTs can also be labeled by microinjection of fluorescently labeled tubulin (Waterman‐Storer, 2002). This method requires a microinjection apparatus and can be more time consuming and technically demanding. Generation of a Stable Cell Line Expressing GFP‐Tubulin
We used a standard method for generating stable NIH‐3T3 cell lines expressing GFP‐tubulin (Sullivan and Satchwell, 2000). Cells are transfected with a commercially available enhanced GFP‐‐tubulin vector containing a neomycin‐resistance cassette (BD Biosciences Clontech).
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Addition of neomycin or G418 (a neomycin analog) selects for cells that incorporate the vector. Resistant clones are screened for GFP‐‐tubulin expression levels and checked for growth rates, migration, centrosome reorientation, and formation of stable MTs (Cook et al., 1998; Gomes et al., 2005). Materials NIH 3T3 cells (ATCC) DMEM (Invitrogen) Enhanced GFP‐‐tubulin mammalian expressing vector (Clontech) Lipofectamine (Invitrogen) G418 (Calbiochem) Cloning cylinders (Sigma) NIH 3T3 fibroblasts are cultured in DMEM supplemented with 10% calf serum (Cook et al., 1998; Gundersen et al., 1994) and kept at subconfluent densities to prevent spontaneous transformation. Transfect cells with GFP‐‐tubulin vector using lipofectamine according to the protocol provided by the manufacture. Two days after transfection, plate cells at low density with DMEM medium supplemented with 1 mg/ml G418. Once colonies are visible by direct observation of the plates (5–7 days after transfection), select individual colonies using cloning cylinders and transfer them to larger plates for expansion. Reduce G418 concentration to 0.5 mg/ml at this stage. When clones have been expanded, check for GFP‐‐tubulin expression by plating cells on acid‐washed glass coverslips, fixing in 20 methanol, rehydrating, and observing by fluorescence microscopy. Select clones with fluorescent MTs that are in a normal radial arrays (Fig. 1B,C). Characterize positive clones by growth rate, migration into the wound, and centrosome reorientation (Fig. 2). We also check clones for formation of stabilized MTs, because this is another rearrangement of MTs that occurs in cells in response to wounding (Cook et al., 1998; Gundersen and Bulinski, 1988). Select clones that behave similarly to parental cells. The level of GFP‐‐tubulin relative to endogenous ‐tubulin is <5% in the selected clone (3T3‐GFPTub) (Fig. 2). Multiple clones should be checked for use in cell assays to ensure that the behavior being studied is typical.
Wound Healing Assay
Wound healing assays are useful to study cell polarization and migration. When a confluent monolayer of cells is wounded, the cells at the wound edge synchronously polarize and begin migrating in the direction
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FIG. 2. Characterization of the NIH 3T3 cell line stably expressing GFP‐‐tubulin (3T3 GFPTub). (A) GFP‐tubulin fluorescence in 3T3‐GFPTub cells. (B) Higher magnification of the inset in (A). The different focal plane shows the MTs of the mitotic spindle. Centrosome reorientation stimulated by LPA or serum is identical in parental NIH 3T3 (C) and 3T3 GFPTub (D) cells. Serum‐starved cells were left in serum‐free DMEM (SF) or treated with LPA or calf serum for 2 h. Cells were fixed with methanol, immunofluorescently stained for centrosomes and MTs, and the extent of centrosome reorientation determined. (E) Expression levels of GFP‐‐tubulin in 3T3‐GFPTub cells determined by Western blot using an anti‐‐tubulin antibody. GFP‐‐tubulin is expressed at <5% the level of endogenous ‐tubulin.
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of the wound. For cells maintained in serum, the polarization and migration responses are triggered as a result of wounding. We introduced a small variation in the wounding assay that allows us to trigger wound response with soluble factors rather than by wounding alone. If confluent monolayers of NIH 3T3 cells are serum starved before wounding, the cells at the edge of the wound do not polarize or migrate until the appropriate serum factors are added (Fig. 3) (Gundersen et al., 1994). The mitogenic lipid lysophosphatidic acid (LPA) is a major serum factor required for MT polarizations in response to wounding (Cook et al., 1998; Palazzo et al., 2001). LPA induces the same MT reorganizations as serum, yet LPA alone is not sufficient to stimulate cell migration (Gomes et al., 2005). Thus, addition of LPA only activates cell polarization and allows the study of polarization events independently from those of active cell migration. To induce both cell polarization and migration responses, serum can be added to the starved cells (Gomes et al., 2005; Gundersen and Bulinski, 1988; Gundersen et al., 1994). Because serum factors are necessary for triggering the responses to wounding, serum starvation also allows a separation of the physical act of wounding from the triggering of polarization and migration responses. This has a number of advantages for studying cell polarization and migration, particularly in studying the very early polarization and migration responses. For example, purified proteins, antibodies, or expression plasmids can be introduced into wound edge cells by microinjection before stimulation, which is not possible with the regular wounding assay.
FIG. 3. Wound assay using serum‐starved monolayers of NIH 3T3 fibroblasts. Schematic representation of a serum‐starved monolayer with MTs, centrosome and nucleus. When the starved monolayer is wounded, centrosome reorientation does not occur. Centrosome reorientation is triggered only on the addition of LPA or serum. (See Palazzo et al. [2001] for details.)
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Introduced proteins can also be tested for sufficiency in induction of cell polarization, which is not possible in a regular wounding assay (Gomes et al., 2005; Palazzo et al., 2001; Wen et al., 2004). We have also found that serum‐starved cells are flatter and provide better subjects for optical imaging. In the following, we describe the specific protocols for serum starvation and wounding of NIH 3T3 fibroblasts. We have found that similar protocols work well for other rodent fibroblast cell lines (Swiss 3T3, Balb 3T3, and NRK fibroblasts). Materials Glass coverslips, washed with 1 N HCl Jeweler’s screwdriver DMEM medium NIH‐3T3 cells Viokase or trypsin/EDTA LPA (1 mM) Preparation of LPA Stock Solution. Add 2.3 ml of a sterile 10 mM HEPES, pH 7.4, 10 mg/ml BSA (fatty acid free, Sigma), 100 mM NaCl solution to 1 mg of LPA (Sigma). Mix well. Aliquot and freeze at 80 . Frozen stocks are good for at least 2 months. Thawed aliquots remain usable for 24 h. Preparation of Starved Cell Monolayers and Wounding. Cover the bottom of a 10‐cm tissue culture dish with coverslips. Detach NIH 3T3 cells from a stock plate using trypsin/EDTA or Viokase and resuspend in DMEM with 10 % calf serum. Dilute cells into 10 ml of DMEM plus 10% calf serum and add 10 ml per 10‐cm dish so that cells will be approximately 20% confluent. Allow cells to grow until just confluent (48 h). Fill three 6‐cm dishes with 8 ml of DMEM without serum. Fill 35‐mm dishes with 1 ml of DMEM without serum (or each well of a 6‐well plate). Using a sterile forceps, transfer each coverslip (containing the confluent monolayer of cells) serially to the 6‐cm dishes containing DMEM. Place the coverslip in the 35‐mm dish and starve the cells for 48 h. Some cells will detach during starvation; however, the remaining cells should still form a confluent monolayer and are fully viable (Gundersen et al., 1994). Using a sterile jeweler’s screwdriver (tip width 1–2 mm), wound the monolayer with an even, continuous movement. Multiple wounds can be made on a single coverslip. Replace cells in incubator and let recover for 20 min. Treat cells with 2 M LPA to induce centrosome reorientation and MT stabilization or add serum (1–5%) to stimulate migration. Centrosome reorientation, MT stabilization, and cell migration can be assessed with fixed cell
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assays (see Cook et al. [1998]; Palazzo et al. [2001]; Wen et al. [2004]), or 3T3‐GFPTub cells can be used for live cell imaging (see the following). Note Wound edge cells can be microinjected after recovery from the wound. Proteins can be microinjected into the cytoplasm, and DNA can be injected into the nucleus. Typically, we find that DNA expression vectors based on CMV promoters allows for efficient protein expression in 1–2 h after microinjection. Because cells will still polarize and migrate if LPA or serum are added up to 8 h after wounding, this allows proteins to be expressed before triggering wound responses. Imaging Centrosome Reorientation in Living Cells
For real‐time imaging of centrosome reorientation, we use 3T3‐ GFPTub cells prepared as described previously. The preparation of the 3T3‐GFPTub cell monolayers for wounding and serum starvation is similar to the preparation of NIH 3T3 cells for wound healing assays (see preceding) with a few modifications to allow for optimal imaging and maintenance of cells under the microscope. 3T3‐GFPTub cells are plated in specific imaging chambers, and modified medium (‘‘recording medium’’) is used for starvation and for maintaining the cells while imaging. Materials Inverted microscope equipped with epi‐fluorescence (Nikon Eclipse TE300) 60 Plan Apo (NA 1.4) objective 40 Plan Fluor (NA 0.6) phase objective Computer‐controlled shutters (Sutter and Uniblitz) on the epi‐illumination light path and the bright‐field illumination light path (for phase‐ contrast imaging) Neutral density filters (ND4 and ND8) on the epi‐illumination light path GFP‐optimized filter set (Chroma) Temperature‐controlled chamber surrounding the microscope Motorized Z‐focus (Nikon) Motorized XY stage (Prior) Cooled CCD camera (Coolsnap HQ)
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Computer with software to control shutters, time‐lapse acquisition, Z‐focus, and XY stage (Metamorph, Universal Imaging Corporation) Recording medium Glass coverslip chambers Preparation of Recording Medium. HBSS (Gibco) containing essential and nonessential MEM amino acids (Gibco), 2.5 g/l D‐glucose, 2 mM glutamine, 1 mM sodium pyruvate, and 10 mM HEPES, pH 7.4. Preparation of Glass Coverslip Imaging Chambers. 35‐mm plastic tissue culture dishes are used. A hole (10–15 mm diameter) is punched in the bottom of the dish with a hand punch tool (Roper Whitney). An acid‐ washed glass coverslip (22 mm2 ) is glued to the bottom of the dish, covering the hole, using Sylgard 184 Silicone Elastomer (Dow Corning) or VALAP (Vaseline/lanolin/paraffin ¼ 1:1:1). Plates are sterilized under UV light overnight. We have also had good results with commercially prepared glass coverslip dishes (35‐mm dish with 10‐mm hole size) from Mattek. Live Cell Imaging of Centrosome Reorientation. Plate 3T3‐GFPTub cells in imaging chambers at a 20% density (see preceding). Once cells reach confluency (after 48 h), wash the cells twice with recording medium and add 4 ml of recording medium. Starve cells in recording medium for 24 h. Equilibrate the microscope temperature chamber at 34–35 for at least 1 h before recording. Wound the monolayer and allow to recover for 20 min in the incubator. Transfer the imaging chamber to the microscope stage. To minimize photodamage (see following), insert the highest neutral density filter in the epi‐illumination light path that still allows observation of the centrosome. Select cells with distinct centrosome and MTs. Minimize as much as possible the exposure of cells to epi‐fluorescence illumination. Determine the minimum exposure time necessary for obtaining an image with a distinct centrosome (see following). Start image acquisition at a rate that does not induce photobleaching (see following) to characterize centrosome position before stimulation. To induce centrosome reorientation, add 100 l of 80 M LPA drop‐wise (2 M final concentration) or 40 l of serum (1% final concentration). Restart image acquisition. We have successfully followed centrosome position up to 4 h after addition of LPA, acquiring 1 image per minute, or up to 18 h after addition of serum, acquiring 1 image every 10 min. By use of this protocol, we were able to observe that centrosome reorientation occurs by nuclear movement away from the leading edge, whereas the centrosome remains at the cell centroid (Fig. 4). If additional information about the nucleus or the cell boundaries are required (for example, to relate centrosome reorientation to leading edge
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FIG. 4. High‐magnification imaging of centrosome and nucleus position during LPA‐ stimulated centrosome reorientation. Selected frames from a time‐lapse movie of a serum‐ starved 3T3‐GFPTub cell before (A) and after (C) the addition of LPA at 0:26 (h:min). The centrosome is identified as a bright point at the center of the MT array. The wound is at the top of the panel. The nucleus is the nonfluorescent oval area. For each time‐point, the position of the centrosome and the centroid of the nucleus were determined and plotted before (B) and after (D) the addition of LPA. Before the addition of LPA, both centrosome and LPA remain in the same position. After the addition of LPA, the nucleus moves away from the leading edge, whereas the centrosome remains stationary at the cell center. Thus, the rearward movement of the nucleus reorients the centrosome. Images were acquired with a 60 Plan Apo (NA 1.4) objective, ND8 density filter, camera binning of 2 2, and 1500 msec exposure at a rate of 1 frame/min. Previously published in Cell (Gomes et al., 2005).
structures, such as lamellipodia or ruffles) sequential phase and fluorescent images can be acquired. To do this, the microscope needs to be equipped with phase or differential interference contrast (DIC) objectives and a shutter in the transmitted light path. The phase or DIC image also contains information regarding possible rotation of the nucleus by reference to the position of the nucleolus and other structures within the nucleus. Metamorph and other commercially available software can be programmed to acquire both epifluorescence and phase images at each time point.
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Notes Chamber Temperature. Preheating the microscope incubator and increasing room temperature minimizes focal drift. We use a temperature of 34–35 instead of 37 , because it is easier to maintain this temperature and focal drift is less of a problem. Recording Media. DMEM medium is not suitable for fluorescence microscopy because it contains riboflavin that interferes with GFP fluorescence microscopy. Riboflavin has an excitation peak at 450 nm and an emission peak at 530 nm, which is similar to enhanced GFP excitation and emission peaks (480 nm and 530 nm, respectively). We also leave out the phenol red that is commonly included in media, because it can absorb light, and this is important for fluorochromes such as rhodamine or DsRed. The recording medium is buffered with HEPES, not bicarbonate, because most microscope chambers are not equipped with a CO2‐enriched atmosphere. The imaging chamber must be covered, because evaporation is significant at 37 , especially during long recordings. Starvation. Starvation time is reduced to 24 h when recording medium is used, because cells start to detach from the coverslip if they are starved for longer times in recording medium. Alternately, cells can be starved with DMEM for 48 h, wounded, transferred to the microscope, and then stimulated with recording medium containing LPA or serum. Starved cells are sensitive to changes in medium; therefore, if the starvation medium is replaced by recording medium without LPA or serum, cells can detaching rapidly (15–30 min). Imaging of the centrosome position before stimulation is not possible if cells are starved in DMEM. Photodamage. Long exposure times can result in photodamage of the cells without detectable photobleaching of the GFP‐tubulin. Photodamage is recognized when cells bleb, MTs depolymerize or, more subtly, do not respond to LPA or serum addition. To minimize photodamage, reduce light exposure as much as possible while searching for cells and during imaging. It is helpful to dark‐adapt one’s eyes by waiting 5 min in complete darkness before searching for cells under the microscope. Also, we have found that the use of neutral density filters in combination with longer exposure times generates less photodamage than short exposure times without neutral density filters. Image Acquisition. Imaging centrosome reorientation can involve long (up to 4 h) recording times. To prevent photobleaching and photodamage, we acquired images at a rate between 1 frame per 1 min and 1 frame every 10 min. These acquisition rates are sufficient to characterize both centrosome and nucleus movements. The exposure used for
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image acquisition should be as short as possible without compromising the visualization of the centrosome. Setting the camera binning to 2 2, acquisition time can be shortened; however, image resolution is also reduced. Triggering Centrosome Reorientation. Centrosome reorientation can be triggered either by LPA or serum, as described previously. The concentration of serum should not be higher that 1%, because serum contains fluorescent factors that increase the background fluorescence, decreasing image quality. This amount of serum is sufficient to induce cell polarization and subsequent migration (Fig. 5). Objectives. We find that a 60 oil Plan Apo objective (NA 1.4) gives the best combination of resolution, sensitivity, and field size for imaging centrosome reorientation. With this objective, the centrosome appears distinct, and individual MTs can be observed (Fig. 4). With a higher magnification objective (e.g., 100), only part of the cell is observed, and photobleaching occurs faster. When only the position of the centrosome
FIG. 5. Low‐magnification imaging of centrosome and nucleus position during serum‐ stimulated centrosome reorientation. Selected frames from a time‐lapse movie of a starved 3T3‐GFPTub cell after the addition of calf serum at 0:00 (min:sec). In this example, a 40 phase objective was used, and both phase‐contrast (top) and epifluorescence (bottom) images were acquired. Although it is not possible to observe individual microtubules in the fluorescence image, the centrosome is visible. The position of the centrosome is represented by a white triangle in the phase‐contrast image. Images were acquired with a 40 Plan Fluor (NA 0.6) phase objective, ND8 density filter, camera binning of 1 1, and 500 msec exposure time at a rate of 1 frame every 3 min.
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is to be determined, we obtained good results using a 40 Plan Fluor objective NA 0.6 (Fig. 5). One advantage of using a lower magnification objective is that higher numbers of cells can be imaged. XY Motorized Stage. Using a motorized XY stage, we have been able to acquire multiple fields (up to 60) at each time point (one frame every 3 min), increasing the speed of data acquisition. A motorized Z‐focus and an autofocus controller are necessary for the acquisition of in‐focus images. Autofocusing can be software controlled. The autofocus software routine must be performed using phase‐contrast illumination to prevent photobleaching of GFP; therefore, a shutter in the transmitted illumination path and phase condenser and objectives are required. We find that 40 dry objectives perform optimally in this scenario. Oil immersion objectives can also be used as long as the travel distances between each field are minimized (<1 cm).
Acknowledgments We thank Jan Schmoranzer for comments on the manuscript. This work was supported by a postdoctoral fellowship from Fundac¸ a˜ o para a Cieˆ ncia e Tecnologia, Portugal (to E. R. G.), and an NIH grant GM062938 (to G. G. G.).
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