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NEMATODE MODULE MBL EMBRYOLOGY

VIEWS: 23 PAGES: 31

									NEMATODE MODULE
MBL EMBRYOLOGY 2009


Faculty:                                         Teaching Assistants:
David Fitch1                                     Elliott Hagedorn4
Paul S. Maddox2                                  David Matus4
Amy S. Maddox2
Joel Rothman3
David Sherwood4
1
  Department of Biology, New York University, Main Building, Room 1009, 100 Washington Square East
 Greenwich Village, New York, NY 10003
2
  Institute for Research in Immunology and Cancer, Department of Pathology and Cell Biology, P.O. Box
 6128, Station Centre-Ville, Montreal, QC H3C 3J7, Canada
3
  Department of Molecular, Cellular and Developmental Biology, University of California, Santa Barbara, CA
93106
4
  Department of Biology, Box 90338, Duke University, Durham, NC 27708



I.      Organization of the Lab

II.     Introduction to C. elegans
        A. Overview
        B. Establishment of embryonic polarity in C. elegans
        C. Vulval induction in the larva
        D. Male mating behavior
        E. Ovulation
        F. Basement Membranes
        G. Comparative development: Natural variation in species related to C. elegans

III.    Techniques Used in this Laboratory
        A. Basic Worm Handling
              1. Picking worms
              2. Mounting embryos/larvae for Nomarski and fluorescence microscopy
              3. Obtaining early embryos
        B. Laser Cell Ablation and Fusion
              1. Cell ablations
              2. Cell fusions
        C. Green Fluorescent Protein (GFP) Markers
        D. RNAi by ingestion of E. coli expressing dsRNA
IV. Experiments
      A. RNAi screen
            1. Sensitized RNAi screen for GEFs that regulate anchor cell invasion

      B. Ablations/Fusions
             1. Ablations to examine cell lineage, cell-signaling, inductions, etc.
             2. Vulval & Gonad Development in C. elegans & Different Species

      C. Male mating behavior
            1. View male mating
            2. View Sensory neurons by labeling with DiI
            3. View male mating in pkd-2 mutant
            4. View male mating in different species

      D. Wild Nematodes
      E. Live cell imaging
            1. Transmitted light imaging
            2. GFP-fusion proteins during the first cell division
            3. View tissue specific marker strains
            4. Observing male tale development in different species
            5. Live analysis of sperm behavior
            6. Ovulation time-lapse
            7. Using photoconversion to track basement membrane dynamics

V. Lab Schedule

VI.   Appendices
      A. Strain List
            1. Mutant strains
            2. Other nematodes
            3. Strains expressing tissue-specific GFP markers
      B. Founder cell lineage and cell fate markers
      C. Sensory organs of the male tail
      D. Gamete maturation and oöcyte ovulation



VII. References




                                                                                      2
I.     Organization of the Lab

We have six formal lab periods beginning on Thursday afternoon. During the orientation period,
we will review techniques that you will need to master in order to perform your experiments
(manipulating worms; mounting embryos, etc.). You will be introduced to a number of major
experimental procedures commonly used in C. elegans embryology laboratories: RNA
interference (RNAi), laser cell ablation/fusion, expression analysis by GFP and ectopic gene
expression. We have available much more than can readily be done by one student and a number
of new experiments for which the outcome is unknown. Keeping this in mind as you read
through this manual, choose experiments you are the most interested in pursuing. The three
David’s, Joel, Elliott, Amy, and Paul will be happy to assist you with experiments in order for
you to become supreme handlers of “the worm”!



II. Introduction to C. elegans

A. Overview

C. elegans, a free-living soil nematode, was chosen by Sydney Brenner in 1963 as an
experimental organism because it is transparent, easy to propagate and is amenable to genetic
manipulation. The entire cell lineage of C. elegans, from the zygote to the adult, has been
determined. Exactly 1090 somatic nuclei are produced during its development to adulthood; 131
cells undergo programmed cell death; adults contain 959 somatic nuclei. Its transparency and
reproducible anatomy make it possible to identify each cell at all stages of development; the
lineage provides us with the knowledge of the ancestry and ultimate fate of every cell. These
features, coupled with its amenability to forward genetic, reverse genetic and molecular studies,
have allowed developmental, physiological and behavioral events to be identified and
characterized at an unprecedented level. The scientific contributions of this small worm are
highlighted by the Nobel Prize for Physiology or Medicine award in 2002 to Sydney Brenner,
Robert Horvitz and John Sulston for their characterization of organ development and
programmed cell death in C. elegans, the 2005 award to Andrew Fire and Craig Mello for their
characterization of RNA-mediated interference, or protein depletion and the 2008 Nobel Prize in
Chemistry to Martin Chalfie for his use of the green fluorescence protein (GFP) as an
experimental reagent in C. elegans.
        C. elegans exists as two sexes: self-fertilizing hermaphrodites, which produce both
sperm and oöcytes, and males. Self-fertilizing hermaphrodites make it possible to propagate
large numbers of animals without the necessity of mating with males. Thus, many worms can be
generated by placing a single hermaphrodite on a petri dish with sufficient food (worms are
grown on a lawn of E. coli). Genetic crosses are performed by mating males with
hermaphrodites; in such a mating, the sperm from the males are preferentially used and the
hermaphrodite serves the role of a female.
        In this lab module, you will have the opportunity to observe the development of wild-
type worms and mutants defective in different aspects of development and behavior. You will
also carry out laser ablation (and/or fusion) of identified cells, analysis of differentiated cell
types by green fluorescent protein (GFP) markers, determination of cell lineages, and reverse


                                                                                                3
genetics by RNAi. One of the main advantages of C. elegans for genetic studies is its short
generation time (3 days). Although the constraints of a short module will not allow sufficient
time to perform genetic experiments, you will have the chance to set up a mating and observe
mating behavior.


B. Establishment of embryonic polarity in C. elegans

The first cell division in Caenorhabditis elegans is asymmetric. Fertilization not only instigates
the completion of maternal meiosis, but also supplies the cell with a single pair of centrioles,
which separate to form two spindle poles shortly after sperm entry. These two events precede the
dynamic movement of cortical cytoplasm from the posterior towards the anterior, and central
cytoplasm moving from the anterior end towards the sperm pronucleus in the posterior. These
cortical and central cytoplasmic flows occur for about 7 minutes, roughly at the same time that
cell fate determinants, such as the posteriorly positioned germ line P granules, become localized
to the posterior. Disruption of the actin cytoskeleton interrupts polarized cytoplasmic flow and
the contents of the first cell are not asymmetrically localized. A group of genes known as the
partitioning defective (par) genes are required for asymmetric division of the zygote.


C. Vulval induction in the larva

Vulval induction in C. elegans has provided an important model for examining cell fate
specification. The entire vulva is derived from a single 1o and two 2o-fated vulval precursor cells
(VPCs), which lie along the ventral epidermis. The 1o and 2o VPCs give rise to lineages of eight
and seven cells, respectively, which have distinctive division and gene expression patterns.
Specification of these VPCs to a vulval fate is dependent on LIN-3, a protein similar to
mammalian epidermal growth factor. LIN-3 is produced by the anchor cell (AC), a specialized
cell in the developing gonad, during the late L2 to early L3 larval stage. LIN-3 activates a
receptor tyrosine kinase pathway in the nearest VPC, P6.p, specifying it to adopt a 1o fate. One
aspect of the 1o fate specification is the upregulation of three distinct ligands for the C. elegans
Notch receptor LIN-12. This upregulation directs the neighboring VPCs, P5.p and P7.p, to adopt
the 2 o-fate. Without the inductive LIN-3 signal from the AC, no vulval induction occurs.
Instead all VPCs divide once and then adopt a non-vulval 3o fate and contribute to the external
epithelium that covers the animal. The resulting vulva-less phenotype can be used as a genetic
tool: Conditional disruption of expression of genes required for vulval specification blocks
vulval formation, and hermaphrodites lacking a vulva cannot lay embryos. Thus, any embryos
that are produced hatch inside the mother and eat the body, causing an easily-scored “bagging”
phenotype.


D. Male Mating Behavior

 C. elegans is an outstanding model system for behavioral analysis. The simple well-
characterized nervous system, complete genome sequence, short generation time and amenability



                                                                                                  4
to genetic analysis facilitate detailed study of behavior at the organismal, cellular and molecular
levels.

Male mating is a complex behavior in C. elegans, which can be divided into the following simple
steps:

1) Response and backing: When the posterior part of a male comes in contact with a
hermaphrodite, the male responds by placing the ventral surface of his tail against her body and
moves backwards, searching for the vulva.
2) Turning behavior: If the male reaches the end of the hermaphrodite (either head or tail), he
will turn and continue to back along the other side.
3) Vulval location: In most cases, the male will stop backing when he scans over the vulva.
4) Sperm insertion: The male inserts his spicules into the hermaphrodite.
5) Sperm transfer: The male then transfers his sperm into the hermaphrodite.



response                           turning                          vulval location




spicule insertion




Figure 2.1 Male mating behaviour



E. Ovulation
During oöcyte maturation in C. elegans, the oöcytes align along the proximal-distal axis of the
gonadal sheath and mature in an assembly-line manner as they proceed proximally toward the
spermatheca (see appendix). Ovulation is the exit of the most proximal oöcyte from the gonad
arm into the spermatheca. This process requires the contraction of the six proximal myoepithelial
sheath cells and the dilation of the distal spermatheca. The sheath contractions pull the dilating
distal spermatheca over the oöcyte, placing the oöcyte into the lumen of the spermatheca where
fertilization occurs. The fertilized egg then moves from the spermathecal lumen to the uterine
lumen through the spermathecal-uterine valve. Fertilization triggers the exocytic events that
promote eggshell formation, and after this point, the fertilized egg takes on its characteristic




                                                                                                 5
oblong “egg shape,” but before this, the large oocyte is remarkably deformable, bounded only by
a plasma membrane, and squeezes through the tiny valves.

F. Basement Membranes

One of the defining structures present in metazoans is basement membrane. This specialized
form of extracellular matrix is thin (60-120 nm), dense and highly cross-linked. Its mechanical
strength provides the structural underpinning for all epithelia, endothelia and many mesenchymal
cells. Basement membranes are also important for organizing tissues into distinct compartments,
tissue repair and in guiding migrating cells during development. About 50 proteins are known to
make up the basement membrane. The main components of basement membranes include type
IV collagen, laminin and nidogen. The major basement membrane components in vertebrates
are all present in C. elegans. Furthermore, in C. elegans these proteins are encoded by much
smaller gene families. For example, laminins are heterotrimeric proteins composed of a
α, β, and γ chain. C. elegans have two α chains , and a single β and γ chain, forming just two
laminin isoforms compared with 15 in vertebrates. Thus, C. elegans is particularly amenable to
genetic analysis of basement membrane formation and function.

G. Comparative development: variation among species related to C. elegans

An important question concerning a "model system" is to what degree are the characters
(features) actually representative of other species. Beyond this, however, variation can be used
to help understand how characters have evolved, how evolving systems adapt to environmental
situations, and what elements of developmental systems are constrained from changing or can
change without deleterious effect to the system. Comparative studies are best done in the context
of a phylogeny, which allows one to infer the direction and number of evolutionary changes. C.
elegans is one of many related free-living species ("rhabditids") that differ to varying degrees in
terms of many characters. For example, some species have a vulva that is positioned in the
midbody; others have posterior vulvae. Many of those with a midbody vulva (like C. elegans)
require an induction signal from the gonad to produce the vulva from ventral Pn.p cells; many
with a posterior vulva are induction-independent. Early cellular events during embryogenesis
also vary substantially. The male tail varies in terms of positions of the rays, the shapes of the
spicules and fan, and the shape of the tail tip, all characters that have been classically important
for taxonomy and phylogenetics. Thus, rhabditids can be used as a powerful system to study
both how developmental mechanisms underlying a conserved feature can change as well as how
development changes to make different forms.


III.   Techniques Used in this Laboratory

A. Basic worm handling

1. Picking up worms. Flame the platinum wire of your pick until it glows red and let it cool
briefly in the air. Put a glob of bacteria on the end of the wire (you may find it helpful to take it
from a worm-free plate, e.g., the one to which you will be transferring worms). Touch the glob
to a worm, using the sticky bacteria to pick up the worm. (Don't "scoop" the worm; just touch the


                                                                                                   6
flattened underside of the pick to it.) Then set the worm down on a new plate in the thick part of
the E. coli lawn. You should try not to gouge either plate as this will encourage the worms to
burrow into the agar, where they become inaccessible. Start by transferring one worm at a time;
eventually it is possible to carry 10 or more worms on one pick-full. Be patient; it takes practice
to get used to transferring worms. Sometimes worms don't like to come off a pick easily; in this
case, soften the glue glob by moving the loaded pick back and forth gently in the E. coli of the
plate to which you are transferring.

TO DO:           Learn how to identify (1) eggs, (2) L1 larvae, (3) L4 larvae, and (4) adults on your
                 N2 plates. Practice transferring worms from plate to plate with a pick.
         Eggs are oval and easy to identify. Adults are the largest worms on the plates and carry
eggs visible in the middle of the ventral side. (Worms crawl on their side, not their ventral
surface.) L1 larvae are the smallest things crawling on your plates. They are sufficiently small
to fit inside an eggshell, which they recently did. At their youngest, they have no dark coloring,
unlike other larvae and adults. L4 hermaphrodites are best identified by their invaginated
developing vulvae that look like pale half moons on the ventral side in the middle of the worm.
There is a black dot in the half moons (see Figure 2.2 below) in L4 larvae.




                               Figure 2.2 L4 hermaphrodite

        On the video display connected to the Nomarski scope, we will look at oogenesis and
early embryogenesis in an intact adult hermaphrodite. We will point out some cells and
structures you will need to identify for yourselves for laser ablations, and we will cover such
basics as identifying left/right and dorsal/ventral. Feel free to go back and forth between
anatomy and lineage charts.

2. Mounting live embryos or larvae for Nomarski (DIC) microscopy. You will have the
chance to view the development of a newly fertilized zygote and perform laser ablations of cells
using the Nomarski scopes. The following outlines the general steps. The techniques are
discussed in more detail below.

       To prepare a Nomarski slide you need to:
       1. identify the desired stage of embryo, zygote, or worm
       2. transfer it to an agar pad on a slide
       3. add a cover slip
       4. seal with Vaseline or Valap (1:1:1 vaseline, lanolin and paraffin)

TO DO in advance:
     1. Ensure that tubes of agar (and Valap) are thoroughly melted.
     2. Prepare a capillary tube for your mouth pipette for liquid transfer of eggs.




                                                                                                   7
         Pull a capillary over a flame so that the end is about the diameter of an adult worm (we
have also prepared pre-pulled capillaries for you). You want it to be big enough to easily suck
up eggs and larvae, but small enough that it can form a seal with a drop of water. Once you
obtain a capillary to your liking, it can be used for all of your experiments. Attach it to a mouth
pipette.
                            2 layers of tape

                                    molten agar
                                                                                   agar pad




                                                                                4th slide
       guide slides
                              Figure 2. Set-up for preparing agar pads




        Prepare an agar pad by placing a generous drop of molten 5% Noble agar on a plain
microscope slide. This slide should be placed between two guide slides whose surfaces have
been raised slightly with two layers of tape so that the agar pad has the proper thickness (Fig. 2).
Work quickly so the agar does not solidify. Perpendicular to these slides, place a fourth slide
over the agar drop so that a pad is formed. Do this by placing one side of the fourth slide at the
edge of the molten agar, avoiding trapping air bubbles in the agar. Press the top slide down
against the supporting slides to flatten the agar. Wait a couple of minutes for the agar to solidify.
        Once the desired specimens are ready to mount, lift the two slides contacting the agar
away from the guide slides. Carefully tease the two apart by a back and forth horizontal motion
that will leave the agar pad on one slide only. Do not rip the pad as you tease the slides apart.
Try to work steadily now so that the agar pad does not dry. (You can pipette some H2O around
the edges of your pad to keep it moist).
        You want to transfer your embryos in a very small volume of liquid. Using the mouth
pipette, take up M9 or egg medium into the capillary (no more than 1-2 microliters or your
embryos will float away!!). Trim the agar pad to approx. 8 mm x 8 mm with a razor blade. Suck
up your eggs up into the capillary and gently blow them in a small volume of liquid onto a corner
of the trimmed agar pad. Using an eyelash mounted to a toothpick, cluster the embryos by
sliding them to the center of the pad. WORK QUICKLY AND KEEP THE EMBRYOS
MOIST. IF THE WORMS DRY OUT THEY WILL DIE. Using your fancy forceps, gently
cover the embryos with a 22 x 22 mm coverslip and seal with melted Vaseline held in the heat
block (back bench in lab).
        To transfer larvae or adult worms to a slide you first add a drop (2.5µl) of M9 on top of
your agar pad. Collect worms as outlined above with your platinum wire, and then gently move
the glob of bacteria containing your worms side-to-side in the drop of M9. This should free most


                                                                                                   8
of the worms from the bacteria. Using your forceps or fingers, gently cover the embryos with a
22 x 22 mm coverslip and seal with melted Vaseline.


3. Obtaining early embryos. Transfer about 20 adult hermaphrodite worms to a drop of about
400 microliters of M9 in a depression well. (We will not be using it here, but often researchers
use a special egg medium if embryos younger than the 2-cell stage are desired). Take two 25
gauge needles and, using them like scissors, cut the hermaphrodite in half near the vulva. This
will release eggs into the medium. Pulling the cut worms in and out of your mouth pipette will
also help to dislodge embryos. Collect or cluster the embryos of the desired stage either with
your mouth pipette or with an eyelash. (See us for needles if you are having trouble pulling the
right size). Using your mouth pipette, transfer the desired embryos to the agarose pad and cover
with a coverslip. Seal with Vaseline to prevent dehydration.


B. Laser Cell Ablation and Fusion

        Laser microsurgery provides a precise and rapid method for eliminating or fusing cells in
the animal. A pulsed dye laser is arranged as an epi-illuminator, its beam being directed
downward through the objective by way of a semi-silvered mirror. Accurate targeting is
achieved by pre-alignment of the focal point of the laser with a reference reticule in the eyepiece.
        We will divide the class into three groups for demonstration; after this instruction you can
sign up for individual time on the laser. If your scheduled or desired time conflicts with
someone else’s, consider that while one person is ablating, another person can be setting up their
next slide. Normal, rigorous technique would require that you regularly monitor your animals
after ablation. This is not practical given our time and equipment constraints. Also, you should
think about the caveats of laser ablation experiments. How do you know when to ablate a cell?
What is the significance of a negative result? What happens to the cytoplasm and cell membrane
after you ablate a cell and how might this affect (or not affect) communication between cells?

1. Cell Ablations. Careful control of the laser pulse power is necessary when performing cell
ablation. Too little power has no effect, and too much power causes excessive damage, often
bursting the embryo. Generally, the best strategy is to deliver several sub-threshold pulses
within a target area. For most ablations, the beam is directed at the nucleus of the cell to be
killed. Usually, debris can be seen to appear in the nucleus after several successful hits. The
beam is directed at various points throughout the nucleus by moving the stage in the x, y, and z
axes. Often, cells that are markedly damaged after ablation will recover and go on to divide; it is
therefore advisable to observe the ablated cell for some time to ensure that it dies.

2. Cell fusions. Cell fusions can be performed to combine the cytoplasmic contents of
neighboring cells. Cell fusions are performed by directing laser pulses to the apposing
membranes of the cells to be fused. Only a few hits are required to completely fuse two cells;
this procedure requires more practice than cell ablation. In most cases a tetrapolar cleavage
occurs during the subsequent cell cycle, generating four mononucleate cells from the fused
dikaryon, all containing cytoplasm from both of the fused cells.



                                                                                                  9
C. Green Fluorescent Protein (GFP) Markers

The green fluorescent protein (GFP) from the jellyfish Aequorea victoria has proven to be an
extremely useful reporter protein in gene expression constructs, since it is visible in living
animals. So useful, in fact, the 2008 Nobel Prize in Chemistry was awarded to Osamu
Shimomura, Martin Chalfie and Roger Y. Tsien for discovering and developing GFP as a
experimental reagent. GFP fluoresces green when illuminated directly with blue light.
Therefore, gene expression can be assayed directly in living worms. This provides us with the
ability to follow dynamic events during development and eliminates the need for fixation and
antibody or in situ staining procedures. The GFP markers can be used in the laser ablation
experiments as well as for visualizing tissue-specific gene expression patterns. To view GFP,
mount embryos as for Nomarski microscopy and observe under the FITC or a narrow-band
GFP(S65C)-specific fluorescence filter. The strains that contain GFP reporters are listed in
Appendix A.


D. RNAi by Ingestion of E. coli expressing dsRNA

The technique of RNAi (RNA-mediated interference) allows an assessment of a loss-of-function
phenotype from a gene of interest virtually overnight; it has revolutionized studies of gene
function in C. elegans and more recently in other organisms as well. After delivery of double-
stranded RNA (dsRNA) to nematodes, phenotypes can be observed in both the treated animals
and in the progeny. Remarkably, the RNAi effect with many genes is potent even when worms
are simply soaked in a solution of dsRNA, or grown on a strain of E. coli expressing sense and
antisense RNAs (which anneal in vivo to make dsRNA). Such ‘feeding plates’ contain a lawn of
concentrated E. coli HT115 harboring a plasmid in which convergent T7 promoters flank an
inserted cDNA fragment. Transcription of both strands occurs through IPTG induction of a
lacO::T7 RNA polymerase chimera.


IV. Experiments

With the exception of the RNAi experiment, you will have your choice of which experiments
from this list to choose from. There are far too many to do them all. We want you to pick the
ones that sound the most interesting to you. Some experiments are more difficult than others
and you might discuss them with one of us before attempting them. To make your lab time most
productive, you should design your experiments prior to the lab period. This is especially true
for the ablations and free periods. Consider what you may need to prepare in advance. For
example, if you want to examine adult hermaphrodites after RNAi, pick the appropriate staged
worms for the treatment.

   A. RNAi—You will work in pairs on the RNAi experiments



                                                                                            10
       Experiment: Sensitized screen for GEFs that Regulate Anchor Cell Invasion:
       The uterine and vulval tissues of the C. elegans hermaphrodite begin development
       juxtaposed but separated from one another by their respective basement membranes
       (BMs). During the third larval stage, a specialized uterine cell, known as the anchor cell
       (AC), connects the two tissues by invading through both tissues’ BMs and into the vulval
       epithelium. The initial connection between these tissues is necessary for the passage of
       embryos out of the adult hermaphrodite. A defect or failure in AC invasion (i.e. a failure
       to connect the uterine and vulval tissue) results in a protruded vulval (Pvl) phenotype—a
       phenotype that can be easily scored using a standard stereomicroscope, thus allowing
       rapid screening for genes that regulate AC invasion.
           Rho family small GTPases are molecular switches that link signaling events at the cell
       surface to reorganization of the actin cytoskeleton within the cell. Rho GTPases exist in
       two forms: an inactive GDP-form and an active GTP-bound form. The switch between
       active and inactive forms is regulated by molecules known as guanine nucleotide
       exchange factors (GEFs; which facilitate the activation of Rho GTPases) and molecules
       refered to as GTPase activating proteins (GAPs; which inactivate Rho GTPases by
       promoting the exchange of GTP for GDP). It is thought that Rho GTPase have
       overlapping functions and that GEFS and GAPs are what provide specificity in when and
       where the GTPases function.
           It is known that the Rho family GTPases cdc-42, mig-2 and ced-10 are expressed in
       the C. elegans AC and play redundant roles during AC invasion (i.e. the phenotype of a
       double mutant between mig-2 and ced-10 is much worse that either mutant alone). This
       kind of functional redundancy makes it difficult to identify genes in a genetic screen, as
       the phenotype of a single gene will be masked by the redundancy of another gene.
           A recent expression screen identified seven GEF encoding genes that were expressed
       in the AC at the time of invasion. Working in pairs, the class will conduct a RNAi screen
       that will target each of the GTPase in combination with one of the seven GEFs. You will
       look for combinations that increase the severity of the Pvl phenotype compared to the
       RNAi targeting the GTPase by itself. This approach should allow us to identify potential
       interactions between the GTPases and GEFs that function to regulate AC invasion.

       Rho Family GTPases: cdc-42, mig-2 and ced-10

       GEFs Expressed in AC: unc-73, ect-2, tag-218, cgef-1, tag-52, pix-1 and tag-77


Outline:

•   We have a 6-well RNAi feeding plate available for pairs of students to examine four genes
    encoding GEFs.
•   The top two wells of each plate contain your controls. The top left well contains RNAi to
    one of the three Rho family GTPase (either cdc-42, mig-2 or ced-10; this will serve as your
    sensitized background and negative control). The top right well contains RNAi to fos-1a, a
    known major regulator of AC invasion (this will serve as your positive control).
•   The remaining four wells have mixed RNAi targeting your Rho family GTPase plus RNAi
    targeting a gene that encodes a GEF.


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•   Tuesday morning, your TAs plated ~100 arrested L1 stage worms expressing an AC specific
    mCherry marker and laminin::GFP, in each well of your 6 well plate. By Thursday evening
    these worms should reach adulthood and show a
    protruded vulval (Pvl) phenotype, should there be a
    defect in AC invasion.
•   We will start by showing everyone how to identify
    and score a Pvl. Then we would like for you to
    compare your positive and negative controls (top
    two wells of your plates).          Once you are
    comfortable with scoring this phenotype, score the
    remaining four wells and determine if RNAi
    targeting a GEF in addition to your Rho family
    GTPase enhances the prevelance of the Pvl
    phenotype.
•   We have created an excel document for everyone
    to input their scoring data into. For the genes that
    most greatly enhance the phenotype of your
    GTPase, you can then score these worms for a
    defect in AC invasion (earlier in development at
    the mid-to-late L3 stage). To do this, the TAs have
    plated worms on the same RNAi plates ahead of
    time and have stages the worms so that they can be
    scored for AC invasion on Friday. Control worms
    will be available so that you can become familiar
    with scoring AC invasion in wild-type animals prior to analyzing your RNAi treated worms.




                                Protruded Vulval Phenotype (Pvl)

       Schedule:
       (TAs)
       Tuesday AM           Plate L1 arrested larva on RNAi plates

       (Students)
       Thursday             Students score RNAi plates for enhanced Pvl phenotype
       Friday-Sunday        Students can score hits from the screen for defects in AC invasion

       Exercises:


                                                                                            12
              a) Look up your GEF genes targeted by RNAi on Wormbase: Wormbase is an
              integrated database that incorporates comprehensive information about C.
              elegans, including all genomic data, literature, genetics, etc. We will demonstrate
              how to use this database, which you can use to look up your gene, find the protein
              domains/similarities, etc.

              b) Scoring for a protruded vulva (Pvl) phenotype: At the beginning of the lab
              section we will demonstrate how to identify and score an RNAi treatment for a
              Pvl phenotype. Then, using your stereomicroscope, comparing the positive (fos-
              1a RNAi) and negative (either cdc-42, mig-1 or ced-10 RNAi) control wells.
              Once you become familiar with identifying Pvls, proceed to score the remaining
              four wells and determine if any of these RNAi treatments increase (enhance) the
              number of worms that have Pvls.

              c) Scoring RNAi hits for AC invasion: The worms on your 6-well plates carry
              two fluorescence markers: a red one in the AC (PLCδ::mCherry) and a green that
              labels the BMs (laminin::GFP). If a particular RNAi treatment enhances the
              number of Pvls on the plate, we have additional RNAi plates that will allow you
              to repeat the treatment and then mount the worms and use one of the compound
              fluorescence scopes (100x objective) to determine if the enhanced Pvl phenotype
              results from a defect in AC invasion.




B. Ablations/fusions

   1. Ablations to examine cell lineage, cell-signaling, inductions, etc.

   a. Ablation of MS. One example of an interesting ablation is the ablation of MS. This
   prevents formation of the pharynx (since the posterior pharynx is produced by MS and the
   anterior pharynx is induced in AB by MS) and eliminates left/right differences in the AB
   lineage, resulting in dramatic cell lineage alterations. You can use the ceh-22::GFP strain
   for these experiments to follow expression of ceh-22 (restricted to the pharynx).

    b. Laser-induced fusions. Fuse oöcytes to make giant embryos. Fuse blastomeres to ask
    which fate is dominant when two cells are made into a heterokaryon.



     2. Vulval & Gonad Development

     The anchor cell (AC) has a key role in inducing the vulva during the L2 stage, but also has
     a more subtle role in vulval patterning during the early L3 stage



                                                                                              13
    a. Laser ablate the anchor cell (AC) in syIs50 (cdh-3::GFP is expressed in the AC) at early
    L3 stage. Is vulval morphology normal? If you save animals with perturbed or no vulval
    induction what happens to the embryos trapped within the animal?

    b. Laser ablate a distal tip cell at the end of one of the gonad arms during the L2 or early
    L3 larval stage or in the adult stage. Recover the worm and let it grow on a plate overnight.
    What happens to the gonadal arm where the distal tip cell was ablated?

    c. Laser ablate the linker cell in the male. Recover the worm and let it grow on a plate
    overnight. What happens to the males gonad when the linker cell is ablated?

     d. Laser ablate the AC or somatic gonad precursors (Z1 and Z4) in other species: e.g.
     Pelodera strongyloides (which has a midbody vulva) and Teratorhabditis palmarum
     (which has a posterior vulva). Try to recover the worms and let them grow for a couple
     days to adulthood. Which ones require induction from the gonad to make a vulva? How
     might you distinguish between experiments showing independence from induction and
     experiments in which your ablation may not have worked?


C. Male mating behavior

      1. View male mating

      Pick L4 stage males onto an NGM (nematode growth medium) OP50 bacterial seeded
      plate and allow males to develop overnight. Pick L4 unc-31 hermaphrodites onto an
      NGM seeded plate and allow to develop for 3 days (we'll pick these ahead of
      time…..older, paralyzed hermaphrodites are used because males can mate and insert
      spicules into these females more easily). Pick 5, three-day old, unc-31 hermaphrodites
      onto an NGM seeded plate with a small spot of bacteria. Place a single male near the
      hermaphrodites and see if you can identify the different steps in mating.

      2. View sensory neurons by labeling with DiI

      With their exposed external cilia, certain chemosensory and mechanosensory neurons
      take up dye in the environment, allowing these neurons to be specifically labeled. The C.
      elegans male tail has several neurons that can be labeled using DiI. You may also like to
      stain males from other nematode species with DiI; you can then compare the organization
      and morphology of these neurons in different species.

      Protocol:

      a. DiI Stock solution is 2 mg/ml in dimethyl formamide, stored at –20oC in a foil wrapped
      tube.

      b. Dilute the stock 1:200 in M9. Some dye will precipitate when you do this; don't worry
      about it.


                                                                                              14
     c. Put 150µl dye solution in a microtiter well, and use a worm pick to transfer some
     worms into the dye. Incubate 2-3 hours at room temperature.

     d. Use a mouth pipette to transfer the worms to a fresh plate, and let them crawl on a
     bacterial lawn for about 3 hours to destain.

     e. Put worms on pads with 0.2% tricaine; 0.02% tetramisole in M9 or 1 mM azide and
     visualize by fluorescence using the appropriate filter.



     3. View male mating in pkd-2 mutant

     Observe mating in a pkd-2 (polycystin-2) mutant and view the expression pattern of a
     pkd-2::GFP reporter construct.

     pkd-2 encodes a putative channel protein and is the C. elegans homologue of the human
     polycystin-2 gene, a gene mutated frequently in polycystic kidney disease.       Is the
     reporter construct expressed in hermaphrodites? Is there anything curious about the
     male expression pattern? In what step(s) in mating are pkd- 2 mutants defective?

     4. View male mating in different species

     Observe mating on plates using a dissecting microscope. What mating positions are
     adopted by the different species? Are there preferences for a particular orientation (e.g.
     parallel versus antiparallel, left coiling versus right coiling)? Map the mating positions
     onto the phylogeny to determine which state is ancestral versus derived. What factors
     may contribute to determining mating position? (Can you make any phylogenetic
     correlations?)

     Can you distinguish between pre- and post-zygotic reproductive barriers between
     species? You could set up interspecific mating tests and determine if mating occurs, if
     hybrid eggs are laid, and if viable F1s are produced. However, be certain you pick
     unfertilized virgins for your experiments. (If a species is a self-fertile hermaphrodite,
     mate them only after they have exhausted all their sperm.)


D. Wild Nematodes

     Nematodes can be collected from the wild outdoors by taking an empty plate and
     sprinkling a small amount of soil from outside onto a fresh agarose plate with E. coli.
     After putting soil on the plate, replace the lid, and place the plate on your bench. As soon
     as a few hours after putting the soil into the plate, Nematodes can be seen crawling on the
     lawn of E. coli. You should transfer these worms to fresh plates. Next, you can collect
     and mount embryos to compare early cell divisions. Collect and mount early embryos as


                                                                                              15
      you would with C. elegans to do timelapse recording of the first cell divisions. Are the
      first cell divisions asymmetric, as with C. elegans? If they are symmetric, how do you
      think localization of P granules occurs? When is the germline specified? What simple
      experiments could be done to determine when P granules are segregated and when the
      germline is specified? The ablation and mating behavior experiments suggested above
      can also be performed on these species.
          Caenorhabditis species are not really "soil" nematodes, but have been found in
      different kinds of habitats. Elegans-group species, such as C. elegans, C. briggsae,
      C. remanei, and some as yet unnamed Caenorhabditis species have been found in
      compost heaps, with reproductive adults found on freshly rotting fruit. (Maybe they
      should be called "fruit nematodes"!) Other Caenorhabditis species have been found in
      association with rotting cactus (C. drosophilae, C. sonorae), dens of burrowing bugs (C.
      japonica) and in phoretic associations (generally as dauer larvae) with snails, pill bugs
      (Armadillidium) and millipedes. If you are interested in trying to discover new
      Caenorhabditis species (there should be many more out there), you could try setting bait
      "traps" with fresh fruit near a compost pile or other place; after a few days, there will be
      nematodes on your now rotting fruit. Check them for distinguishing characteristics of
      Caenorhabditis or Elegans-group species (e.g. prominent median bulb in the pharynx, fat
      ray 6 in the male tail, "hook" structure on the precloacal sensillum).


E. Live cell imaging

      1. Transmitted light imaging. Many important events in the early embryo can be
      observed by transmitted light imaging. The cytoplasm is largely filled with refractile yolk
      granules, which bear witness to the flows that occur during the establishment of anterior-
      posterior polarity. The cortical cytoplasm flows towards the anterior, and the central
      cytoplasm flows towards the posterior. The nuclei are clearly visible as round clearings in
      the granular cytoplasm. The sperm pronucleus, initially residing in and defining the
      posterior, expands from 2-4 microns compacted to 7-10 microns at maximum. This
      characteristic increase in size is an indication of cell cycle progression. Nuclear envelope
      breakdown is visible. The centrosomes, or microtubule organizing centers of the mitotic
      spindle, are also visible as clearings in the cytoplasm, and their increase in size is also
      characteristic of mitotic progression. It is possible to see yolk granules and other features
      aligned along the long straight microtubules (themselves invisible) that emanate from the
      centrosomes towards the cortex.

      One main strength of using the C. elegans early embryo as a model system for cell
      biology is the reproducibility of timing, placement, magnitude, etc of events. You can
      perform time-lapse imaging of several embryos and measure a developmental event and
      compare the timing, etc. among your specimens.

      2. GFP-fusion proteins during the first cell division. Using the method described
      above for dissecting embryos for live cell imaging, isolate embryos from the appropriate
      transgenic strain. (For example, observation of chromosome dynamics requires TH32,
      GFP:Histone H2B and GFP:gamma tubulin). We will also have fluorescent markers for
      components of the cortical actomyosin cytoskeleton. Image the embryos by DIC and


                                                                                                16
fluorescence using widefield or spinning disk confocal microscopy. Record several
untreated (control) divisions prior to timelapse recording sequences from RNAi treated
worms to ensure a well-controlled interpretation. See the microscopy handbook for
details on setting up the microscopes and proper alignment. In untreated embryos, note
the timing and rates of nuclear migration, nuclear envelope break-down (NEBD), spindle
orientation, anaphase onset, cytokinesis onset and completion, as well as the relative sizes
of the resulting blastomeres.

We will have available RNAs directed against genes whose products are implicated in
cell division. You could treat L4 stage worms with soaking RNAi to deplete target
proteins from the gonad and resulting embryos. Compare cell division parameters in
control and RNA-treated embryos and devise a quantitative method to characterize the
role of the RNAi target protein in the first cell division.


3. View tissue specific marker strains. In the appendix is a list of transgenic strains
with GFP expressed in different tissues. These strains are available at the front of the lab,
and be easily mounted and viewed for fluorescence.


4. Observing male tale development. Pick early L4 males and mount them onto agar
pads with levamisole anesthetic. Watch morphogenesis occur over a period of 2-4 hours.
This can also be done with other species.

To observe how cell boundaries change during male tail morphogenesis (e.g. cell fusions
in tail tip and seam cells), use a him-5 (high incidence of males) strain that has ajm-1::gfp
(adherens junction molecule-1, a component of belt adherens junctions in epithelia).

5. Live analysis of sperm behavior. Mate red (histone) sperm males with
hermaphrodites of other strain
Use mCherry::Histone males.
Observe the migratory behavior of the red sperm in the spermatheca
If the hermaphrodite had green sperm (eg strain TH32), what happens to the red histones
after fertilization? Are thered histones retained in the male pronucleus or displaced by
green histones from the “mother” wrom?When is the red histone gene from the sperm-
derived chromosome transcribed, translated, and incorporated into chromosomes in
blastomeres?

6. Ovulation time-lapse. A time-lapse study of ovulation in C. elegans can be
performed by first anesthetizing adult hermaphrodites in a solution of M9 with 0.2%
tricaine and 0.02% levamisole for 30 minutes in a glass depression slide. The
anesthetized worms are then mounted on 5% agarose pads with 10-20 µl of anesthetic
and covered with an 18 mm glass coverslip. Seal the edges of the coverslip with
Vaseline and capture images at 10-20 second time intervals by DIC optics using a 100X
objective. Try performing time-lapse by also visualizing the sperm with Cyto dye above



                                                                                          17
or sperm containing red-fluorescent protein-tagged histone H2B, which incorporates into
the DNA.

7. Using photo-conversion to study basement membrane dynamics. Recently, the
photo-switchable fluorophore Dendra2 was developed for use in C. elegans. Based on
principles similar to photo-activatable GFP where a light induced conformational change
in the fluorophore alters the fluorescent properties of the molecule, Dendra2 initially
fluoresces under blue light (green emission, just like GFP), however, after brief exposure
to UV light Dendra2 converts and is thereafter excited by green light (red emission).
Using the laminin::Dendra2 transgenic worms and an appropriate UV laser, you should
be able to photo-convert a region of interest within the basement membrane (for example,
under the anchor cell prior to invasion) and then track the fate of the basement membrane
that was converted from green to red fluorescence. Alternatively, you could photo-
convert all of the basement membrane in the animal from green to red fluorescence and
then image the animal at later time points to determine how newly synthesized laminin is
added to a growing basement membrane (the new laminin will be green, whereas
preexisting laminin will remain red).

8. Vital dye staining of other species. We will have several vital dyes available for you
to try out, including mitotracker, ER-tracker, and Lyso-tracker from Molecular Probes.
See Dave Matus for the protocol to visualize these fluorescent markers.




                                                                                       18
VI. Lab schedule

Lab 1, Thursday Afternoon

      General Demo Introduction (All Faculty and TAs)
      Set up nematode mating plates

             Individual demos:
                     a) mounting early embryos (Paul/ Amy)
                     b) mounting and dismounting larvae (Elliott)
                     c) laser ablation and fusion (Dave S)
                     d) wormbase (Dave M)

Lab 2, Thursday Evening

      Score RNAi plates for enhanced Pvl phenotype
      Chalk Talk: Amy Maddox
      Overview of non-RNAi experiments (All faculty)
      Independent experiments

Lab 3, Friday Afternoon

      Score RNAi screen hits for defects in anchor cell invasion
      Independent experiments


Lab 4, Friday Evening

      Score RNAi screen hits for defects in anchor cell invasion
      Independent experiments
      Friday Evening Lecture Series, 8 o'clock, Lillie Auditorium


Lab 5, Saturday afternoon

      Score RNAi screen hits for defects in anchor cell invasion
      Independent experiments


Lab 6, Saturday evening

      Chalk Talk: Paul Maddox
      Independent experiments




                                                                    19
V. Appendices

A. Strain list (transgenes are annotated as promoter::PROTEIN)
      N2 (Wild-type Caenorhabditis elegans var. Bristol)
      Caenorhabditis briggsae
      Caenorhabditis remanei
      Oscheius tipluae
      Oscheius dolichouroides
      Teratorhabditis palmarum
      Rhabditis myriophila
      Rhabditis dudichi
      Pristonchus pacificus
      Panogrolaimus sp. PS1159
      Panagrolaimus superbus
      Acrobeloides butschlii
      Cephalobus sp. (DWF1301)
      Zeldia sp. (DWF1701)
      Panagrellus redivivus (MT8872)
      Mesorhabditis miotki
      Pelodera strongyloides
      Pellioditis typical
      Mesorhabditis longespiculosa
      ajm-1::GFP
      ceh-22::GFP
      elt-2::GFP
      edIs20
      lin-26::GFP
      hlh-1::GFP
      cdh-3::GFP
      cog-2::GFP
      unc-31
      him-5
      pkd-2::GFP
      pkd-2; him-5
      pAC::mCherry, laminin-beta::GFP
      laminin::GFP
      laminin::Dendra2
      nmy-2:: NMY-2::GFP (JJ1473)
      pie-1::GFP::tubulin (AZ235)
      pie-1::GFP::H2B (AZ212)
      pie-1::GFP::PLCδPH (OD58)
      pie-1::GFP::UNC-59 (OD26)
      GFP::gamma-tubulin; GFP::Histone (TH32)
      OD122
      F57B10.1>GFP::beta-tubulin
      cdh-3>beta-tubulin::GFP


                                                                 20
1. Mutant strains

him-5. High incidence of male strain. Approximately 16% of the self-progeny are male.

pkd-2. pkd-2 encodes an ortholog of human PKD2 (OMIM:173910; mutated in
autosomal dominant polycystic kidney disease) that is expressed in the cilia of three types
of male-specific sensory neurons.


2. Strains expressing tissue-specific GFP markers

Strains listed below contain integrated transgenes that direct tissue-specific expression of
the Green Fluorescent Protein (GFP). Most of the GFP transgene strains we have
brought are homozygous for a chromosomally integrated ‘S65C’ variant reporter and are
marked by a dominant ‘roller’ phenotype (animals have a “C” shaped appearance).
Depending upon the nature of the fusion transgene, fluorescence may be cytoplasmic
(e.g. edIs20), localized to subcellular structures (e.g. ajm-1::GFP).


ajm-1::GFP expresses in epithelial adherens junctions, thus outlining all epithelia. This
can be used to study cell fusion, cell movements, etc. ++ (adults); + (embryos and
larvae).

ceh-22::GFP: expresses in differentiated pharynx muscle cells. +++ (all stages). Pharynx
cells are made autonomously from MS and by induction in AB; thus, this marker is useful
for detecting early embryonic interactions.

elt-2::GFP (rrIs1): expressed in E lineage (~4E) and differentiated gut cells (very strong
fluorescence, visible under the fluorescence dissecting microscope). This strain does not
carry the Rol marker. +++ (all stages)

edIs20: This strain carries a fusion of a putative signal transduction gene (F25B3.3) to
GFP. Expression begins in embryonic neural precursors and is later seen in a large part
of the nervous system. In particular, the ventral nerve cord and commissural interneuron
processes are most impressive. The edIs20 strain is in a Him genetic background,
meaning that it generates spontaneous males at high frequency due to X-chromosome
nondisjunction. The male mating structure, the bursa, contains sensory rays that can be
visualized with this marker (very strong fluorescence, visible under the fluorescence
dissecting microscope). +++ (late embryos and older)

lin-26::GFP: expresses in nuclei of hypodermal and other non-neuronal ectodermal cells.
+ (embryos); ++ (later stages)

hlh-1::GFP: expresses in body wall muscle cells. ++ (all stages)


                                                                                         21
       cdh-3::GFP: (syIs50) high levels of expression in the anchor cell and seam cells
       cog-2::GFP: Transcriptional reporter for the cog-2 gene, which encodes a sox domain
       protein expressed in the uterine pi cells that form the mature uterine-vulval connection.
       cog-2 is also expressed in body wall muscles.

       nmy-2::GFP: Non-muscle myosin chain. Expressed in the germline and somatic gonad.

       pkd-2::GFP: (nIs128) pkd-2 encodes the C. elegans homologue of the human disease
       gene polycistin-2. pkd-2::GFP is expressed in the male-specific sensory neurons of the
       head (the CEMs), rays and hook (HOB).

       Beta-tubulin strains:
       F57B10.1>GFP::beta-tubulin expresses GFP::ß-tubulin under the control of 2kb of the
       F57B10.1 promoter. It is expressed in a wide variety of tissues during larval development
       and can be used to visualize microtubules.

       cdh-3>GFP::ß-tubulin can be used to visualize microtubules in the Anchor Cell(AC)
       from the L3 stage and in the HSN neuron in the L4 stage.


       Embryonic and germline GFPs

       The pie-1 promoter drives expression in the germline: the gonad and embryos. This
       fairly strong promoter is routinely used with GFP fusions to study protein behavior in the
       early embryo in lieu of identifying and isolating the protein of interest’s own regulatory
       sequences.

       pie-1::tubulin::GFP: Expressed in germline and early embryos.

       pie-1::GFP::H2B: Histone::GFP fusion in germline

       OD3 pie-1::tubulin::GFP: Expressed in germline and early embryos.

       AZ244 pie-1::GFP::H2B and pie-1::tubulin::GFP: in germline

       TH32 pie-1::GFP::H2B and pie-1::gamma-tubulin::GFP: in germline


B. Founder cell lineages and cell fate markers

         Consider the following lineage when you design your ablation experiments. The general
experimental outline is to ablate a cell or subset of cells, and then assay the fates of the progeny
of the remaining cells. A variety of fate markers are available. However, we suggest using one
of the GFP-expressing strains. This will allow you to directly assess the results of your ablation
after allowing time for development.


                                                                                                 22
                                            Po
                                                               P1
                                                   EMS                     P2
                                                                     P3



Founder cell               AB                MS          E          P4    D      C

Gut cells                                                20

Seam cells!!!!!!! !20!!!!!!!!!!!

Pharynx : structure        16                  2
             muscle        19                 18

Body muscle                 1                 28                          20     32

Embryonic deaths           98                 14                                 1


     GFP expressing strains. The constructs that cause tissue-specific GFP expression are
     particularly useful markers (see the strain list).
     Gut granules. The cytoplasm of intestinal cells contains refractile and autofluorescent
     "gut granules" ("Rhabditin"). In intact embryos they begin to appear at about the 100-
     150 cell stage (3-4 hours post fertilization). These can be observed under polarized light.
     Simple polarized light can be obtained by carefully pulling out the Wollaston prism from
     the Nomarski scope. Gut nuclei can be visualized with elt-2::GFP.
     Pharynx morphology. Look for the distinct structures of the pharynx tissue. Some
     ablations may produce a partial pharynx, such as the posterior bulb. You may want to
     refer to pharynx morphology of an intact L1. Pharynx muscle can also be visualized with
     ceh-22::GFP.
     Muscle twitching. Early in the morphogenesis stage of wild type development (6-7
     hours post fertilization), "twitching" movements start. These are indicative of muscle
     function, and the embryo soon develops into a squirming larva. If body muscles are
     present in partial embryos, they can form functional tissues, and may begin twitching at
     about the same developmental time. This activity will subside over time. Muscle cells
     can also be visualized with hlh::GFP.




                                                                                             23
C. Sensory organs of the male tail




(From the worm atlas)

Sensory Rays
 9 bilateral pairs of sensory rays, numbered 1 (anterior) to 9 (posterior). Each ray has a structural
cell and two sensory neurons (RnA and RnB). Except for ray 6, all neurons have contact with
the outside. The sensory rays are required for response to hermaphrodites dorsally and play a
role in the male's ventral response to hermaphrodites in combination with ventrally located
sensory structures, e.g. hook sensillum, hook and spicules. Rays are also involved in turning
behavior.

The Hook Sensillum
The hook sensillum is composed of 2 sensory neurons (HOA and HOB) and two support cells,
and is closely associated with the hook, a sclerotic structure that is shaped like a hook and
derived from a single cell. This entire structure can be removed by ablation of the epidermal
P10.p cell in the late L2 or early L3 stage. The hook sensillum is required for vulval location.

The Post-cloacal Sensilla (p.c.s)
Each post-cloacal sensilla is made up of 3 sensory neurons--PCA, PCB and PCC--and three
support cells. These sensilla may be involved with precisely positioning the spicule over the
vulval opening.

The Spicules
Each spicule contains 2 sensory neurons, SPD and SPV. The processes of these neurons extend
the length of the spicule and are open to the environment. Each spicule also has a motor neuron,
the SPC. The spicule structure is, not surprisingly, required for sperm transfer. The SPD sensory


                                                                                                  24
neuron and SPC motor neuron are also required for spicule insertion. The SPV sensory neuron
regulates sperm transfer.

D. Gamete maturation and oöcyte ovulation




from: McCarter et al., Developmental Biology 205, 111-128 (1999)




                                                                                        25
VII. References

General:

htpp://wormbook.org

Bowerman, B. (1998). Maternal control of pattern formation in early Caenorhabiditis elegans
     embryos. Curr. Topics in Dev. Biol. 39, 73-117.

Bowerman, B. (1999). Maternal control of polarity and patterning during embryogenesis in the
     nematode Caenorhabditis elegans. In Cell Lineage and Cell Fate Determination, Acad.
     Press, pp. 97-117.

Brenner, S. (1974). The genetics of Caenorhabditis elegans. Genetics 77, 71-94.

Dolinski, C., J.G. Baldwin and W.K. Thomas (2001). Comparative survey of early
       embryogenesis of Secernentea (Nematoda) with phylogenetic implications. Canadian
       Journal of Zoology 79, 82-94.

Hird, S.N. and J.G. White, Cortical and Cytoplasmic Flow Polarity in Early Cells of
       Caenorhabditis elegans. Journal of Cell Biology, 1993. 121(6): p. 1343-1355.

Kimble, J. and D. Hirsh (1979). The postembryonic cell lineages of the hermaphrodite and male
      gonads in Caenorhabditis elegans. Dev Biol 70, 396-417.

Labouesse, M, and Mango, SE (1999). Patterning the C. elegans embryo: moving beyond the
      cell lineage. Trends Genet 15, 307-13.

Newman-Smith, E. and Rothman, J.H. (1998). The maternal-to-zygotic transition in embryonic
     patterning of C. elegans embryos. Curr. Op. Genetics Dev. 8, 472-480.

Seydoux, G., and Fire, A. (1994). Soma-germline asymmetry in the distributions of embryonic
      RNAs in Caenorhabditis elegans. Development 120, 2823-2834.

Sternberg, P.W. (2001). Working in the Post-Genomic C. elegans World. Cell 105, 173-176.

Sulston, J.E. and H.R. Horvitz (1977). Post-embryonic cell lineages of the nematode,
       Caenorhabditis elegans. Dev Biol 56, 110-56.

Sulston, J.E., et al. (1983). The embryonic cell lineage of the nematode Caenorhabditis elegans.
       Dev Biol 100, 64-119.




                                                                                              26
Early Development:

Cell-Autonomous Development


Bowerman, B., et al. (1993). The maternal gene skn-1 encodes a protein that is distributed
     unequally in early C. elegans embryos. Cell 74, 443-52.


Boyd L., et al. (1996). PAR-2 is asymmetrically distributed and promotes association of P
      granules and PAR-1 with the cortex in C. elegans embryos. Development 122, 3075-
      3084.

Cowan, A.E. and J.R. McIntosh (1985). Mapping the distribution of differentiation potential for
      intestine, muscle, and hypodermis during early development in Caenorhabditis elegans.
      Cell 41, 923-32.

Doe CQ, Bowerman B. (2001) Asymmetric cell division: fly neuroblast meets worm zygote.
     Curr Opin Cell Biol. 13, 68-75

Draper, B.W., et al. (1996). MEX-3 is a KH domain protein that regulates blastomere identity in
       early C. elegans embryos. Cell 87, 205-216.

Etemad-Moghadam B., et al (1995). Asymmetrically distributed PAR-3 protein contributes to
      cell polarity and spindle alignment in early C. elegans embryos. Cell 83, 743-752.

Goldstein, B. (1996). Specification of the anteroposterior axis in Caenorhabditis elegans.
       Development 122, 1467-1474.

Guo, Su, and Kenneth J. Kemphues (1995). par-1, a gene required for establishing polarity in C.
      elegans embryos, encodes a putative Ser/Thr kinase that is asymmetrically distributed.
      Cell 81, 611-620.

Hunter, C.P., and Kenyon C. (1996). Spatial and temporal controls target pal-1 blastomere-
       specification activity to a single blastomere lineage in C. elegans embryos. Cell 87, 217-
       226.

Kemphues, K.J., et al. (1988). Identification of genes required for cytoplasmic localization in
     early C. elegans embryos. Cell 52, 311-20.

Kemphues, K. (2000). PARsing embryonic polarity. Cell 101, 345-348.

Lin, R., Thompson, S., and J.R. Priess (1995). pop-1 encodes an HMG box protein required for
        the specification of a mesoderm precursor in early C. elegans embryos. Cell 83, 599-609.




                                                                                              27
Maduro, M.F. and Rothman, J.H. (2002). Making Worm Guts: The Gene Regulatory Network of
      the C. elegans Endoderm. Dev Biol 246, 68-85.

Mello, C.C., et al. (1992). The pie-1 and mex-1 genes and maternal control of blastomere identity
       in early C. elegans embryos. Cell 70, 163-76.

Nance, J., and Priess, J.R. (2002). Cell polarity and gastrulation in C. elegans. Development.
       129, 387-97.

O'Connell, K.F. (2000). The centrosome of the early C. elegans embryo: inheritance,
      assembly, replication, and developmental roles. Curr Top Dev Biol 49, 365-84.

Wallenfang, M.R. and Seydoux, G. (2000). Polarization of the anterior-posterior axis of C.
      elegans is a microtubule-directed process. Nature 408, 89-92.

Zhu, J., Hill, R.J., Fukyama, M., Sugimoto, A., Priess, J.R., and Rothman, J.H. end-1 encodes an
        apparent GATA factor that specifies the endoderm precursor in Caenorhabditis elegans
        embryos. Genes Dev. 11, 2883-2896.

Zhu, J., Fukushige, T., McGhee, JD, and Rothman, JH. Reprogramming of early embryonic
       blastomeres into endodermal progenitors by a Caenorhabditis elegans GATA factor.
       Genes Dev. 12, 3809-3814.



Cellular Interactions:

Evans, T.C., et al. (1994). Translational control of maternal glp-1 mRNA establishes an
       asymmetry in the C. elegans embryo. Cell 77, 183-94.


Goldstein, B. (1992). Induction of gut in Caenorhabditis elegans embryos. Nature 357, 255-7.


Hutter, H., and R. Schnabel (1994). glp-1 and inductions establishing embryonic axes in C.
       elegans. Development 120, 2051-2064.


Mello, C.C., B.W. Draper, and J.R. Priess (1994). The maternal genes apx-1 and glp-1 and
       establishment of dorsal-ventral polarity in the early C. elegans embryo. Cell 77, 95-106.

Meneghini, MD, Ishitani, T, Carger, JC, Hisamoto, N, Ninonmiya-Tsuji, J, Thorpe, CJ, Hamill,
     DR, Matsumoto, K, and Bowerman, B. (1999). MAP kinase and Wnt pathways converge
     to downregulate an HMG-domain repressor in Caenorhabditis elegans. Nature 399:793-
     7.



                                                                                               28
Mickey K.M., et al. (1996). An inductive interaction in 4-cell stage C. elegans embryos
      involves APX-1 expression in the signaling cell. Development 122, 1791-1798.


Priess, J.R., H. Schnabel, and R. Schnabel (1987). The glp-1 locus and cellular interactions in
        early C. elegans embryos. Cell 51, 601-11.

Priess, J.R. and J.N. Thomson (1987). Cellular interactions in early C. elegans embryos. Cell 48,
        241-50.


Rocheleau, C.E., Downs, W.D., Lin R., Wittmann C., Bei Y., Cha Y.H., Ali M., Priess J.R. and
      Mello, C.C. (1997). Wnt signaling and an APC-related gene specify endoderm in early
      C. elegans embryos. Cell 90, 707-716.

Rocheleau, CE, Yasuda, J, Shin TH, Lin, R, Sawa, H, Okano, H, Priess, JR, Davis, RJ, Mello,
      CC (1999). WRM-1 activates the LIT-1 protein kinase to transduce anterior/posterior
      polarity signals in C. elegans. Cell 97, 717-726.

Schelesinger, A, Shelton, CA, Maloof, JN, Meneghini, M, and Bowerman, B. (1999). Wnt
       pathway components orient a mitotic spindle in the early Caenorhabditis elegans embryo
       without requiring gene transcription in the responding cell. Genes Dev 13:2028-38.

Thorpe, C.J., Schlesinger, A., Carter, J.C., and Bowerman, B. (1997). Wnt signaling polarizes an
      early C. elegans blastomere to distinguish endoderm from mesoderm. Cell 90: 695-705.

Thorpe CJ, Schlesinger A, Bowerman B. (2000). Wnt signalling in Caenorhabditis elegans:
      regulating repressors and polarizing the cytoskeleton. Trends Cell Biol 10:10-7


Shin, TH, Yasuda, J., Rocheleau, CE, Lin, R, Soto, M., Bei, Y, and Davis, R.J. (1999). MOM-4,
       a MAP kinase kinase kinase-related protein, activates WRM-1/LIT-1 kinase to transduce
       anterior/posterior polarity signals in C. elegans. Mol Cell 4: 275-80.



Male Mating Behaviour:

Barr, M. M., J. DeModena, et al. (2001). “The Caenorhabditis elegans autosomal dominant
       polycystic kidney disease gene homologs lov-1 and pkd-2 act in the same pathway.” Curr.
       Biol. 11: 1341-1346.

Barr, M. M. and P. W. Sternberg (1999). “A polycystic kidney-disease gene homologue required
       for male mating behavior in C. elegans.” Nature 401: 386-389.




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Garcia, L. R., P. Mehta, et al. (2001). “Regulation of distinct muscle behaviors controls the C.
       elegans male's copulatory spicules during mating.” Cell 107: 777-788.

Hedgecock, E. M., J. G. Culotti, et al. (1985). “Axonal guidance mutants of Caenorhabditis
      elegans identified by filling sensory neurons with fluorescein dyes.” Dev. Biol. 111: 158-
      170.

Liu, K. S. and P. W. Sternberg (1995). “Sensory regulation of male mating behavior in
      Caenorhabditis elegans.” Neuron 14: 79-89.

Simon, J. and P. W. Sternberg (2002). “Evidence of a mate-finding cue in the hermaphrodite
      nematode Caenorhabditis elegans.” Proc. Natl. Acad. Sci. 99: 1598-1603.

Sulston, J. E., D. G. Albertson, et al. (1980). “The Caenorhabditis elegans male: Postembryonic
       development of nongonadal structures.” Dev. Biol. 78: 542-576.



Ovulation:

Bui, Y.K., and Sternberg, P.W. (2002). "Caenorhabditis elegans inositol 5-phosphatase homolog
negatively regulates inositol 1,4,5-triphosphate signaling in ovulation." Mol. Biol. Cell. 13:
1641-1651.

Clandin, T.R., DeModena, J.A. and Sternberg, P.W. (1998). "Inositol trisphosphate mediates a
RAS-independent response to LET-23 receptor tyrosine kinase activation in C. elegans." Cell.
92: 523-533.

McCarter, J., Bartlett, B., Dang, T., and Schedl, T. (1999). "On the control of oöcyte meiotic
maturation and ovulation in Caenorhabditis elegans." Dev. Biol. 205: 111-128.

Maddox, A.S., B. Habermann, A. Desai, and K. Oegema. (2005) Distinct roles for two C.
elegans anillins in the gonad and early embryo. Development. 132: 2837-48.

Wolke U, Jezuit EA, Priess JR. (2007) Actin-dependent cytoplasmic streaming in C. elegans
oogenesis. Development. 134: 2227-36.


Uterine and vulval development:

Blelloch, R. and J. Kimble (1999). “Control of organ shape by a secreted metalloprotease in the
       nematode Caenorhabditis elegans.” Nature 399: 586-90.

Hanna-Rose, W. and M. Han (1999). “COG-2, a Sox domain protein necessary for establishing a
      functional vulval-uterine connection in Caenorhabditis elegans.” Development 126: 169-
      179.


                                                                                                 30
Kimble, J. (1981). “Alterations in cell lineage following laser ablation of cells in the somatic
      gonad of Caenorhabditis elegans.” Dev. Biol. 87: 286-300.

Kimble, J. and D. Hirsh (1979). “The postembryonic cell lineages of the hermaphroidite and
      male gonads in Caenorhabditis elegans.” Dev. Biol. 70: 396-417.

Newman, A. P., J. G. White, et al. (1996). “Morphogenesis of the C. elegans hermaphrodite
     uterus.” Development 122: 3617-3626.

Wang, M. and P. W. Sternberg (2001). “Pattern formation during C. elegans vulval induction.”
      Curr. Top. in Dev. Biol. 51: 189-220.

Sherwood, D. R., and Sternberg, P.W. (2003). Anchor cell invasion into the vulval epithelium in
      C. elegans. Dev. Cell 5, 21-31.

Sherwood, D. R., and Sternberg, P.W. (2005). Fos-1 promotes basement membrane removal
      during anchor cell invasion in C. elegans. Cell In press.

Sommer, R.J. and Sternberg, P.W. (1994). Changes of induction and competence during the
     evolution of vulva development in nematodes Science 265, 114-118.

Sommer, R.J., Carta, L.K. and Sterberg, P.W. (1994).         The evolution of cell lineage in
     nematodes. Development Suppl., 85-95.

Sommer, R.J. and Sternberg P.W. (1996). Evolution of nematode vulval fate patterning. Dev
     Biol., 173: 396-407.

Van Buskirk, C.V. and Sternberg, P.W. (2007). Epidermal growth factor signaling induces
     behavioral quiescence in Caenorhabditis elegans. Nature Neuroscience, 10: 1300-1307


Basement Membranes:

Kramer, J.M. Basement membranes. WormBook. 2005 Sep 1:1-15




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