Cell migration is a basic developmental function that serves to build tissues, organs, and whole animals. Defects in cell migration are associated with birth defects and cancer, in particular the metastasis of tumors. Over the past forty years researchers have used the fruit fly to understand the genetic basis of development, including cell migration, but many of the tools and approaches used are beyond the skills and understanding of an undergraduate and advanced high school lab. We have developed a practical lab that allows students to use fly oogenesis to understand how genes regulate cell migration. Students learn to sort males from females, recognize fly genetic markers to identify wild type and mutant animals, hand-dissect ovaries, perform histochemical staining to reveal gene expression in this tissue, and visualize normal and aberrant cell migration using light microscopy to distinguish the effect of a key mutation in a gene required for cell migration. From this approach, students learn how mutations can aid in understanding gene function and how modern genetic tools and microscopy are used to study gene expression and development. Because these genes have human homologs, students learn how model organisms can be used to understand the molecular basis of disease and disorders, such as cancer.
Cell biological and genetic approaches have revealed that in organisms as diverse as worms and humans, cells migrate by turning on and off genes, first to loosen connections with their neighbors, then to change shape and flatten to crawl along precise extracellular matrix paths. Migrating cells are able to recognize their proper destinations and stop to build the detailed structure of a tissue (Aman & Piotrowski, 2010).
In humans, mutations in the genes that control cell migration are associated with birth defects and cancer. Cancer is a complex disease with many defects in cell metabolism, cell proliferation, cell death, and the kind of uncontrolled cell invasion characteristic of tumor metastasis. Metastatic cancer cells change their adhesivity, leave a primary site and invade surrounding tissues, often resulting in patient mortality. These aberrant cell behaviors mimic normal cell functions during development, but the identity and function of the genes in these processes are poorly understood (Naora & Montell, 2005).
Over the past forty years, genetic and biochemical approaches have revealed many of the genes controlling cell migration (Rorth, 2007; Rorth, 2009). These genes encode proteins that function at all levels of the process: outside the cell to modify the extracellular matrix, on the surface of the cell to change its adherence or “stickiness,” in the cytoplasm to remodel cell shape, allowing the cell to squeeze through its surroundings, and in the nucleus to turn on and off the genes that regulate the process. Many of these genes have been identified using a model organism, in particular genes regulating the cell cycle in yeast and Caenorhabditis elegans (Edwards, 1999; Norbury & Nurse, 1990)
Here we have developed a lab that allows undergraduates and advanced high school students—really, a student at any level with some dexterity and patience—to visualize the effect of a mutation in the slow border cells gene (abbreviated slbo, where the gene is italicized and lower case, and the protein it encodes is upper case, Slbo) in the migration of the border cells during Drosophila oogenesis (Montell et al., 1992). Slbo (pronounced “slow-bo”) is homologous to the human transcription factor CCAAT/enhancer binding protein (C/EBP) that acts in the nucleus to turn on and off genes required for migration. Human C/EBP is associated tumor formation and tumor suppression, depending on the tissue examined (Nerlov, 2007). Using a stock of flies that carries the slbo mutation and are readily available from the Indiana University Drosophila Stock center in Bloomington (http://flystocks.bio.indiana.edu/) and with minimal set up, students can sort mutant and wild type animals, dissect and stain ovaries, and mount the tissue on slides to visualize normal and defective cell migration. From this exercise, students learn how genetics can be used to study normal development and to understand the genetic basis of cancer.
Drosophila melanogaster is the premiere genetic organism to study the role of genes in developing tissues. Fruit flies are easily reared and maintained, and stocks carrying mutations or transgenes can be ordered from stock centers around the world. You need to know several basic things about Drosophila to carry out these experiments.
Learning about Gene Expression Using the slbo Enhancer Trap
Flies have useful genetic tools that allow scientists to study the function of their genes and where these genes are turned on or “expressed” in tissue. Transposable elements are “jumping genes” that are engineered by cutting and pasting DNA, and then injecting it into fly embryos, where the DNA will jump into the fly chromosome to establish stocks bearing the transposon in every cell of the transgenic organism (Stanford et al., 2001). The transposon we will use is called an “enhancer trap” and is built to contain two useful genes: (1) an eye color marker gene, which gives the animal red eyes to confirm that the transposon is inserted in the fly chromosome; and (2) a reporter gene used to see the expression of a gene that the enhancer trap has “hopped” next to. A commonly used reporter gene is lacZ, which produces the protein β-galactosidase. β-galactosidase can be seen in fixed tissue by the histochemical stain, X-gal, which is colorless until it is cleaved by β-galactosidase to yield the products galactose and 5-bromo-4-chloro-3-hydroxyindole, which is then oxidized into 5,5'-dibromo-4,4'-dichloro-indigo, an insoluble blue product. Where the blue product accumulates is tissue is where the lacZ enhancer trap reporter gene is active.
The fly stock we use here carries a lacZ enhancer trap transposon insertion that has “jumped” next to the gene slbo (Montell et al., 1992). The slbo-lacZ insertion is stable, lacZ will not jump back out, and the enhancer trap insertion does two important things for this lab: (1) it disrupts the slbo DNA sequence and effectively creates a mutant allele (version) of the slbo gene, designated slbo10310; and (2) it carries the lacZ gene, which gets turned on in the same tissue as the slbo gene, revealing a blue color in a pattern identical to the normal slbo gene. Thus migrating cells expressing slbo-lacZ are marked in blue, and the effect of the slbo mutation on the progress of cell migration can be readily detected.
Learning about Fly Genetics Using the slbo Mutation
The slbo gene is on the second chromosome. The slbo fly stock carries two homologous chromosomes. One has the enhancer trap mutation inserted next to the slbo gene. The other has no insertion, has a wild-type slbo gene, meaning that it has a normally functioning copy of slbo, and carries a curly-wing mutation elsewhere on the chromosome. This curly-marked chromosome is a special “balancer chromosome”—yet another useful genetic tool in flies—(1) that carries a curly-wing marker that allows it to be recognized; (2) that includes multiple inversions that prevent recombination; and (3) that is lethal in two copies. When students dump out and sort flies from the slbo stock, they will be able to recognize heterozygous, curly-winged flies (genetically written slbo10310/CyO) and homozygous, straight-winged flies (genetically written slbo10310/slbo10310) (Figures 1 and 2). (The CyO/CyO animals are embryonic lethal and so will not be seen as adults.) The real strength of this lab is that homozygous mutant animals are plentiful in this stock, allowing student to dissect and see the mutant migration phenotype.
Learning about Cell Migration Using the slbo Marker Gene
In this lab, students dissect separately the ovaries from wild-type (curly-wing) and mutant (straight-wing) females (Figures 1, 2, 3), and stain the respective tissue for slbo-lacZ gene activity. slbo-lacZ becomes active during mid-oogenesis when 16–20 border cells break off, “delaminate,” from the anterior of the egg chamber and migrate as a cluster through the nurse cells to arrive at the border between nurse cells and oocyte (Figures 1 and 4). In a light microscope, slbo-expressing cells appear blue in stained tissue from the slbo enhancer trap stock, and reveal the progress of this migration both in normal slbo/CyO and in mutant slbo/slbo ovaries. Mutant slbo/slbo ovaries fail to finish border cell migration and fail to form properly a micropyle in the eggshell of the mature egg; as a result, the egg cannot be fertilized, and so the female is infertile (Figure 4).
Learning about the Use of Model Organisms to Understand Human Disease
Results will show both the progress of normal migration and the effect of mutations on this developmental event. From these observations, students will appreciate the power of model organisms to give insight into the role of genes in basic developmental processes, with implications for the mechanism of tumor formation. Model organisms have a short generation time, and mutants are easily obtained. Their genomes have been completely sequenced, and most human genes associated with disease are present, allowing functional studies. Certain tissues, such as the ovary used here, allow easy screening for mutations that disturb development. Moreover, the links between genes can be rapidly made in model organisms, so that networks of genes regulating development can be identified. Because these networks are common to humans and model organisms, much can be learned about the function of homologous genes in normal and abnormal tissue growth.
Learning Goals and Objectives
Goal: Determine the requirements for slbo gene function for cell migration by visualizing migration and expression in normal and mutant animals.
Chemicals (all solutions except XGAL can be stored at room temperature)
PBS (phosphate-buffered saline)
PBS + BSA (PBS + a pinch of bovine serum albumin [Sigma Chemicals, St. Louis, MO])
PBT (PBS + 0.1% Tween 20 [Sigma])
FIX (4% paraformaldehyde = ¾ part PBS, ¼ part 16% paraformaldehyde [Polysciences])
Stain solution for 50 mL:
1M Na2HPO4: 0.684 mL (of 1M stock)
1M NaH2PO4: 0.316 mL (of 1M stock)
5M NaCl: 1.5 mL (of a 5M stock)
IM MgCl2: 0.05 mL (of a 1M stock)
K4Fe(II)(CN)6: 0.065 g (yellow crystals)
K3Fe(II)(CN)6: 0.051 g (orange crystals)
Triton X 100: 1.5 mL (of 10% stock)
H20: 47 mL
XGAL (10 mg mL−1 in N'N'-dimethyl formamide, stored at −20°C)
Mounting solution (50% glycerol in PBS)
microfuge tube rotator (Labquake, Thermofisher)
depression well slide
dissection micro forceps (Dumont #5 Forceps)
dissection needle (Fisher 08-965B)
glass Pasteur pipets
slbo10310/CyO flies (Bloomington Stock Center, stock #12227)
vials and stoppers (Genesee Scientific)
food recipe: You can make this food, pour it into vials, and store the vials covered in the fridge for 2–3 weeks (http://flystocks.bio.indiana.edu/Fly_Work/media-recipes/molassesfood.htm).
dry yeast (Fleishman's brand jars and packets)
Flynap and Drosophila anesthetizers (Carolina Biological). Note that new anesthetizers need holes poked in the bottom with a sharp probe.
paintbrushes (Utrech 234 white nylon sable)
Set up (6–8 weeks) requires growing enough flies to satisfy the class size (step 1, below). Conditioning flies requires a five-minute procedure but must be done in two successive days before dissection (step 2). Dissection, staining, and visualization (steps 3–5, Figure 3) require three class periods and need not be done on successive days.
STEP 1: Instructor preparation (begin two months before). The slbo10310 flies can be ordered from the Indiana Stock center (stock #12227; ordering information at http://flystocks.bio.indiana.edu/). Upon arrival, the fly stock is reared at room temperature and expanded by flipping the adults into fresh food vials every 5–10 days and splitting the emerging adults among several vials. Keep the old vials for 30 days maximum; freeze and dispose of plastic vials in the regular trash, or soak, wash and rinse thoroughly glass vials for re-use. In this way, the stock can be expanded significantly over the course of 1–2 months. Expect that a 20-day-old vial of flies should yield sufficient animals for two groups. Planning ahead at this point should ensure that the number of slbo10310 flies will not be a limiting factor in carrying out this lab.
STEP 2: Instructor or student group preparation (begin two days before). Females must be “conditioned” on two successive days on fresh vials containing dry yeast/water mixture. Feeding triggers a physiological cue to lay eggs; thus well-conditioned females will contain ovaries that are easy to dissect and will contain all the stages of oogenesis, so that migration can be followed easily by microscopy. Flip 30–40 adult flies (males and females) from the stocks you have expanded into a fresh vial containing 4–5 drops of water and a sprinkling of dry yeast. Do this again the next day. By the third day when students dissect the ovaries, the organs should be large and easy to dissect.
STEP 3: Dissecting, fixing, and staining ovaries (Lab period 1). What follows is the formal, in-class lab exercise.
Put the slbo10310/CyO flies to sleep in an anesthetizer with a few drops of Flynap. Sort male and female fruit flies under the dissection scope with a paint brush. To distinguish males and females, rely on animal size (smaller males), posterior cuticle banding pattern (solid black on males, striped on females), and the presence of sex combs on males (Figure 2).
Separate the female flies into two groups: one with curly wings (genetically slbo10310/CyO) and one with straight wings (genetically slbo10310/slbo10310) (Figure 2).
Dissect ovaries in the following way (Figure 3):
Place a depression slide under the dissecting scope and fill the well with PBS/BSA.
Transfer a straight-wing female to a dry area next to the depression well on the slide. This is your operating table.
Focusing on the female through the dissection scope, use the forceps to pinch the female under the “armpits” (Figure 3A).
Holding the female in this way, with the other hand, use a dissection needle to tug slightly and remove the posterior-most ovipositor structure (Figure 3B).
Press on the abdomen and force the ovary into the PBS/BSA (Figure 3C).
Look for the ovary pair, and use the forceps and probe to carefully isolate it from other organs (Figure 3D). Even if the organ breaks up at this step, sweep the parts into the depression well.
Repeat on several more straight-winged flies, dissecting 10–15 ovaries into the depression well, and then use a Pasteur pipet coated with PBS/BSA to transfer all the tissue to a microfuge tube. Label the tube “non-CyO,” and repeat the dissection for the CyO genotype.
Wash each tube for 5 minutes with 400 µL 1xPBS, as follows: Add PBS to the microfuge tube, rotate end-over-end for 5 minutes, let the ovaries settle to the bottom, then remove as much PBS as possible without removing the ovaries. Be very careful to watch that you do not suck up any tissue at this step; a good scientist will always save the washes in another container to check for accidental removal of ovaries.
Use gloves to add 300 µL FIX solution to each tube, and rotate for 5 minutes.
Use gloves to carefully remove the FIX from each tube, and wash in 400 µL of PBT three times for 5 minutes each. Let the ovaries settle between washes and carefully remove the supernatent.
Add 400 µL of Embryo Stain Solution (ESS) with XGAL (1ml ESS with 11 µL XGAL) to each tube.
Wrap each tube in aluminum foil and incubate overnight at 37°C with rocking.
STEP 4: Washing and mounting ovaries (Lab period 2)
Remove stain solution from each tube, and wash three times for 5 minutes each in 400 µL PBT.
Break up the ovaries into individual egg chambers by repeatedly pipetting the tissue 3–4 times through a small pipet tip until they break apart and appear as a “snow” of isolated egg chambers. Be careful not to overdo this as the sheer force degrades the tissue. Allow to settle thoroughly, and then remove wash.
Add 400 µL mounting solution and rotate for 5 minutes.
Let ovaries settle another 5 minutes.
Mount tissue on a labeled microscope slide in mounting solution in the following way: Cut thin strips of electrician's tape and affix to center of slide 15 mm apart (a bit less that the width of a coverslip). Then transfer the ovaries to the area between these “feet” and cover them with coverslip. Paint the edges of this sandwich with nail polish to seal and glue coverslip in place. Store in slide box.
STEP 5: Visualizing ovaries (Lab period 3)
Examine the slide under the microscope. Compare and contrast wild-type slbo10310/CyO and slbo10310/slbo10310 mutant egg chambers. Describe and sketch exactly what you see.
Typical Experimental Observations and Results
Students should be able to see normal migration occur in the heterozygous flies, and a block in migration in the homozygous mutant flies (Figure 4). The mutation in slbo reduces gene activity, resulting in a range of migration defects, from none to partial migration, that can be documented by the student.
Questions for the Students
What kind of genes might slbo regulate to drive cell migration?
Why is the enhancer trap insertion in the slbo gene create a weak mutation?. What does that mean? If a strong, knock-out mutation is embryonic lethal, what are the advantages of a weak mutation? Why is migration partial in slbo/slbo ovaries? What are the ranges of migration?
Why is staining stronger in the homozygous mutant?
slbo/slbo animals are not fertile. How could you prove that?
Based on the results you get, what is the next experiment that should be done?
Technical Problems, Preparation of Materials, and Laboratory Safety
Removing ovaries cleanly requires some training, but even a sloppy dissection will yield some tissue that should be kept for staining. The staining step gives reliable results as long as fixation is kept to 5 minutes. Flynap is harmless, and capping tubes can minimize its perfumed odor. FIX should be made fresh; PBS/BSA will go bad over the course of a month, but all other chemicals are good for more than a year, if stored properly. FIX and XGAL should be handled with gloves, but the small amounts used here do not require a fume hood. Drosophila melanogaster are not related to the Mediterranean fruit fly, an agricultural pest, so D. melanogaster can be disposed of in a fly morgue. (A fly morgue is a funnel over a 500-ml beaker with 300ml water and a drop of liquid detergent to break the surface tension. It can be cleaned out regularly by flushing down the sink.)
Evaluation of Student Understanding
Assessment techniques include a quiz before the lab, and a data sheet to document the extent of migration at each stage for wild-type and mutant ovaries observed.
Discussion and Conclusions
This exercise introduces the student to a fly model of cell migration. In a practical sense, the student gains an understanding of genetics and skills in tissue dissection, histochemical staining, and microscopy. From a conceptual standpoint, the student learns the link between gene expression and protein function, and the effect of a mutation on a developmental process. The connection of cell migration to cancer cell metastases can quickly engage students' interest because of the impact of cancer on their own lives. The ability of metastatic cancer cells to detach from a primary tumor site, enter the bloodstream or lymph system, and adhere to secondary sites, leading to inoperable spread of the cancer, provides a jumping-off point to discuss how genes might serve as targets of therapies and how model organisms can be used to understand the genetic basis of human disease.
We have implemented this lab at both college and high school levels with success. Growing a sufficient number of flies requires pre-planning so that students have enough biological material to perfect their dissecting skills; the slbo stock can be maintained indefinitely by flipping it into a fresh vial once a month, or re-ordered (several months in advance) from the Bloomington Indiana Drosophila stock center. The dissection is one of the simplest in biology, and some students quickly become adept at the technique to collect sufficient tissue for subsequent staining steps. The staining is dependable, and as long as no tissue is lost at the washing steps, the defect in migration is conspicuous so that many examples should be visible under the light microscope. Graduates come back and report to us that this lab was one of the highlights of their academic experience.
This lab supports the Next Generation Science Standards (NGSS Lead States, 2013): High School Life Science 1 (HS-LS1), “From Molecules to Organisms: Structures and Processes.” Students who demonstrate understanding of this lab can: (1) “construct an explanation based on evidence for how the structure of DNA determines the structure of proteins which carry out the essential functions of life through systems of specialized cells” (HS-LS1-1), and (2) “develop and use a model to illustrate the hierarchical organization of interacting systems that provide specific functions within multicellular organisms” (HS-LS1-2).
Sexing Drosophila: http://arrogantscientist.wordpress.com/sexing-drosophila/
We thank the students in our labs whose enthusiasm has encouraged us to put together this protocol to disseminate this useful approach to the study of cell migration. We also thank Denise Montell, Pernille Rorth, and Alan Spradling for developing the enhancer trap line and the border cell migration system.