Are there cell-cycle dependent variations in cadherin-mediated cell adhesion?
Maria Cimpean
Department of Natural Sciences, Colby-Sawyer College, New London, NH
ABSTRACT
Cell adhesion makes possible the development of tissues and organs. It has thus been an important topic in biomedical research, cancer research in particular. One method used to quantify cell adhesion, the dual pipette assay, has shown persistent variability in the force required to separate adherent cells despite efforts to minimize variability through strict protocols and assay automation (1). We are using FUCCI (fluorescence ubiquitination cell cycle indicator) to determine if cell-cycle dependent changes might account for the observed variability in separation force. The FUCCI technique involves antiphase oscillating proteins that mark the cell-cycle transitions with corresponding fluorescent probes (2). Comparison of fluorescence distributions in aggregated and unaggregated FUCCI E cad cells showed preferential accumulation in aggregates of cells in the G1 stage of the cell cycle.
BACKGROUND
One of the prerequisites for the evolution of multicellular organisms was the possibility of cells to adhere to one another. At some point, the association of cells rather than their isolation conferred evolutionary advantages. As one may imagine, the subsequent development of tissues and organs depended on regulated and sophisticated cell-cell adhesion mechanisms.
One of the first experiments that laid the groundwork for the cell adhesion research field was Wilson’s. He separated the cells of a live sponge through a fine sieve, thus obtaining mostly single cells. Interestingly, he observed that with time, those individual cells formed agglomerations of cells, and ultimately grew to become whole new sponges. Wilson performed the same experiment with two different species of sponges, and found that the separated cells did not mix with those of another species (3,4). The question then became: how is this possible? What are the features that make it happen?
At least part of the answer involves the properties of the cell surface. It is now known that each type of cell has a different set of proteins on its surface, and some of these differences are involved in the formation of tissues and organs during development. E.E.Just’s (1939) (5) observations suggested that the cell membrane was different in different cell types, and Townes and Holtfreter (1939-1955) had demonstrated that when pieces of amphibian embryonic tissue combined, the result was an anatomically correct structure; Holfreter observed that some tissue pieces associated preferentially, such as mesoderm with endoderm or ectoderm, but not endoderm with ectoderm. He later found that this specific affinity was also present in individual cells. Holfreter placed pieces of amphibian embryonic tissue in an alkaline pH (approximately 10), which caused the tissue to “dissolve” in a suspension of single cells, and observed that when normal pH conditions were restored, even if the cell suspension had been previously mixed/disorganized, the cells are able to reaggregate so that the structure of the embryo is restored (6,7). Moscona (1952, 1961) expanded this experiment to include later stage avian and mammalian embryos, and introducedtrypsinization as a tissue dissociation method.However, the mechanism by which cells actively organize into tissues rather than being randomly sorted was still unknown (7).
In 1964, Steinberg introduced the differential adhesion hypothesis, a model based on the patterns of cell sorting and tissue spreading behavior he observed. He believed that thermodynamic principles explained these patterns, and he suggested that movements within the embryo are an attempt to restore an equilibrium state that was previously disrupted by a change in gene activity, which changed the cell surface molecules. He proposed that cells interact in a way that would form an aggregate using the smallest free energy, and thus the most thermodynamically stable pattern. Steinberg explored this model further, and in 1996, Foty and his colleagues in Steinberg’s laboratoryused a system consisting of cells expressing identical adhesion molecules, but at different levels, and demonstrated that the lower-expression line enveloped the higher-expression line, thus showing that different strengths of adhesion is all that is needed for sorting to occur. The higher-expression line had more “glue” on its surface, so those cells were able to have a higher number of stable contacts/adhesions between each other.The numbers of adhesion molecules, as well as their type and properties (such as strength) are responsible for creating boundaries between tissues, as seen in the embryo with germ layers (4,7).
Three classes of proteins generally make up complexes that represent the functional units of cell adhesion: cell adhesion molecules/adhesion receptors, extracellular matrix proteins, and cytoplasmic plaque/peripheral membrane proteins (8). While there are many classes of molecules involved in the mediation of cell adhesion, cadherins seem to be the major class of adhesion molecules responsible for calcium-dependent cell-cell adhesion in vertebrate tissues; thus, cadherins are critical for the initiation and maintenance of connections between cells. This class of adhesion molecules consists of calcium-dependent adhesion molecules, hence the name cadherins (9).Calcium-dependent adhesion molecules coexist with calcium-independent adhesion molecules, but cross-adhesions occur only between cells expressing the same type of adhesion molecules, as demonstrated by Takeichi in his work with vertebrate cells. Takeichi’s lab later (1981 and after) discovers that cadherins are actually a large superfamily of related cell surface proteins (10).
Cadherins are anchored into cells by catenins,and interact with other cadherins on neighboring cells. The cadherin-catenin complex is involved in the formation of adherens junctions that connect epithelial cells. Cadherins are also well known for their involvement in cell recognition and cell sorting during development (10). One of the members of the cadherin family found in epithelial tissues, and named after the type of tissue it was found in is E-cadherin (also called L-CAM) (11). If E-cadherin is absent from the epithelium, the other cell adhesion and cell junction proteins cannot maintain cell-cell adhesions. In addition to this role, there is evidence that suggests E-cadherin may act to suppress the invasiveness and metastasis of epithelial tumor cells(8).Since cancerous cells need to be able to detach from the primary cancer site and enter circulation and/or invade another tissue, and then proliferate to form new tumors, it is not surprising that any molecule that maintains cell-cell adhesions would prevent this from happening (12). The loss of original tissue contacts involves changes in cell-cell adhesion and cell-extracellular matrix adhesion. Therefore, studying the role of cell adhesion could lead to a better understanding of the development of cancer, and thus lead to a more effective approach to cancer treatment (13, 11).
One of the methods used to quantify cell adhesion, the dual pipette assay, measures the force required to separate a pair of cells. It has been shown that adhesions strengthen over time until a plateau is reached. However, there is some variability in the force measurements for various contact periodsdespite almost complete automation of the process (1). It is assumed that the main (if not only one) source of variability is the cells themselves. The variability might be due to cell health, cell culture conditions, cell density, transfection efficiency, and other potential factors.The stage of the cell division cycle that the cells are in during the contact periods could be a source of variability that would be worth exploring. The simplest and most well-known supporting evidence is the observation that, in culture, cells undergoing mitosis detach from the substrate, “round up”, and divide; after, they reattach to the substrate. It is clear that there is a change in the adhesive properties of cells as they are progressing through the cell cycle. In order to determine if there is indeed a notable change in adhesive properties, and whether this change is significant, we need a method of testing adhesion that would also allow us to know, at any time, which phase of the cell cycle the cells tested are in.
The cell cycle, the universal process by which cells reproduce, is also vital in the development of tissues and organs; it is the process that underlies development of all life. The study of cell cycle commenced with the discovery of cell division. Kolliker’s realization that early embryonic cleavage represented a series of cell divisions was further extended by 1890 by Pringsheim, Strasburger, and Hertwig, who determined that gametes (eggs and sperm) were single cells that joined to eventually form an organism. This showed that even the most complex multicellular organism passed through a single celled stage. More details were observed as microscopes and microscopic techniques improved, and work started in the 1950s eventually led to the discovery of cell cycle engines such as cyclin-dependent kinases (CDKs). Cyclins were discovered as a result of a search for proteins with fluctuating levels throughout the cell cycle, and this discovery led to the identification of CDKs, which have a cyclin subunit and a kinase subunit. CDKs drive cells through the cell cycle, and are regulated by the availability of cyclin subunits (14). For example, cyclin A/cdk2 accumulates during S phase and its activation triggers the transition to G2, a phase characterized by the accumulation of cyclin B/cdc2. The transition from G2 phase to mitosis is triggered by the Cdc25-mediated dephosphorylation of the cyclin B/cdc2 complex (MPF) (15). The activation of cyclin B/cdc2 that is necessary for G/M progression is a fairly well-characterized step of the cell cycle (16).
One of the markers of the transition of a cell from G2 to mitosis is the activation of a protein-serine/threonine kinase, which has been identified as MPF in frog eggs, as histone H1 kinase in sea urchin eggs, and as growth-associated histone H1 kinase in mammalian cells (17). A commercially-available G2 to M transition marker works by monitoring expression levels and localization of the functional elements of cyclin B1 fused to EGFP. This fluorescent reporter moves from the cytoplasm to the nucleus during the M phase (18). Kinases are also involved in cell-cell adhesion, and an inter-dependence between the cell cycle and adhesions has been established (19).
The cell division cycle is a fundamental and complex process that lies at the core of every proliferating cell, so researchers had to design techniques that would allow them to analyze this process. Attaching a fluorescent probe to a molecule that appears only in one stage of the cell cycle is a relatively simple method to identify cells found in that specific stage. Each of the molecules mentioned above could potentially be tagged by attaching fluorescent probes. A fairly recent technique for visualizing cell-cycle transition is the fluorescent ubiquitination-based cell cycle indicator (FUCCI). Developed in 2008, this technique employs an idea similar to the simple tagging technique mentioned above. It involvesantiphase oscillating proteins that mark the cell-cycle transitions and corresponding fluorescent probes. The probes label individual G1 phase nuclei red and those in S/G2/M phases green. The G1/S transition is marked by yellow nuclei, as Cdt1 (tagged with RFP) levels decrease and geminin (tagged with GFP) levels increase. Cdt1 and geminin are regulators of the cell cycle. Thispermits unprecedented spatial and temporal resolution,which makes it possible to better understand how the cell cycle is coordinated with other biological events such as cell-cell adhesion. So far, time-lapse imaging of these cells was used to explore the spatiotemporal patterns of cell-cycle dynamics of cultured cells, and the development of tumors across blood vessels in mice. However, while the Fucci technique allows us to follow cell division in a population by indicating the cell cycle stage, it does not discriminate between the S, G2, and M phases (2, 20).
Figure 1.FUCCI, the new tool to visualize the cell cycle. From: MechaliLutzmann, Cell, 2008
Objective:To explore the potential relationship between different stages of the cell cycle and cell-cell adhesiveness using the FUCCI system.
Hypothesis: Cell adhesiveness varies with the stage of the cell cycle.
The variability of cell-cell adhesiveness observed using the dual pipette assay is at least partly due to the stage of the cell cycle that the cells are found in during the time they are in contact, since any other factors that could arise from the manipulations of the cells have been eliminated. Therefore, the variability has to originate from the cells themselves. It is possible that at least one stage of the cell cycle (M, G1, S, or G2) will exhibit less/more adhesive cells. In other words, my hypothesis is that differential cell-cell adhesiveness will be observed when different stages of the cell cycle are compared.
Null hypothesis:
- There is no significant difference in terms of cell-cell adhesiveness between cells in different/any stage of the cell cycle. (Since cell adhesion is vital in the development of tissues and organs, significant differences are not feasible.)
- The variability of cell-cell adhesiveness observed using the dual pipette assay is due to the genetic variability of the cell population. This might cause (among other factors) cells to have different affinities for certain molecules, which might cause the variability of cell-cell adhesion within the same phase of the cell cycle.
The study design used is schematically represented below:
Figure 2. Study Design.
METHODS
Cell culture
FUCCI cells expressing different levels of mouse E-cadherin were generously provided by Dr. Chu YehShiu. The cells were cultured at 37 °C with 5% CO2, in DMEM (Lonza) supplemented with 10% fetal bovine serum (FBS) (Lonza), 1/50 L-glutamine (Lonza) and 1/100 penicillin/streptomycin (Lonza).
Cell preparation
The cells were kept in Ca2+- containing solution in order to maintain their adhesive properties. Upon aspiration of the medium from the 25 cm2 flask, the cells were rinsed twice with Hepes Buffer Saline Magnesium Free (HMF). The cells were then incubated in 1ml of 0.01% Trypsin (Sigma, porcine pancreas Type IX-S, lyophilized powder, 13,000-20,000BAEE units/mg protein) in HMF solution for 15 min in the CO2 incubator. Afterwards, the cells were dispersed, transferred to a 15 ml centrifuge tube, and centrifuged for 5 minutes at setting #4. The supernatant was then removed and 5 ml of HMF were added. The cells were then dispersed and re-suspended in HMF. After the removal of the HMF solution, 5 ml of 0.01% soybean trypsin inhibitor were added to the pellet of cells. The cells were then dispersed and re-suspended in HMF once again. The supernatant containing 0.01% soybean trypsin inhibitor (Sigma) was removed, followed by the dispersion of the cells in the 2 ml Hepes Buffer Saline Calcium Magnesium Free(HCMF) to effectively disperse for single cells.
Single cells dissociation was verified with the help of a hemocytometer.The dissociation/dispersion should always performed with a fine tipped long bore, fire polished, glass Pasteur pipette. Before the cells were counted, 3ml of HCMF was added to the solution and the cells were dispersed. Then the cells were counted and the volume was adjusted so that the starting concentration is always the same (2*10^6 cells/ml) for all subsequent repetitions of the experiment. Cells in HCMF were then centrifuged and the supernatant is discarded. The required volume of medium is added to pellets of cells in the tube and dispersed. This dissociation is not sterile and the cells are kept on ice throughout the entire process in order to minimize any adhesion formation (21).
Aggregation assay
In order to gather data on the cell cycle stage distribution throughout the aggregates, the cells kept in Ca2+-containing solution (as described above) are dissociated, and then allowed to adhere by performing an aggregation assay. The purpose of this method is to allow individual cells to collide and form aggregates. A 4-well plate (Nunc) containing 0.5 ml of the cell suspension (2 x 106 cells/m in warm medium) in each well is placed on a gyratory shaker set at 90 rpm, and in a 37ºC environment. After set periods of time (e.g. 5 minutes),the aggregates, preferably containing 5-20 cells for ease of visualization, as well as the cells that are not part of the aggregates were visualized under the microscope using fluorescence.
Data/Image acquisition
Imaging was accomplished through the use of a Zeiss Im35 microscope equipped with an objective lens(40X N.A.= 0.75), a Lumen Dynamics X-Cite 200DC illuminator, a CCD camera,and iVision software. Within the iVision software, an exposure of 200 ms and a 2 x 2 bin size was used. Phase and fluorescence microscopy was used. Photos of both the dissociated cells in suspension and the aggregates were acquired and the cells in each of the phases of the cell cycle were counted. In order to visualize the cells and differentiate between the stages on the same image, images of the red and green fluorescence emitted by the cells will be taken.The 2 separate black and white images were pseudo-colored (green and red, depending on the filter used) and then merged. The merged image shows the green, red, and yellow cells, which representscells in all the phases of the cell cycle besides mitosis. The cells in mitosis can be counted by subtracting the total number of pseudo-colored cells from the total numbers of cell counted in a non-fluorescent phase image.
Statistics
Mean and standard deviation were calculated. The chi square test was used to determine whether there is a statistically significant difference between the expected and observed categories.Results were determined to be statistically significant if the calculated p-value was less than the 0.05 significance level.
RESULTS