Methods for studying oogenesis.

2. Methods to manipulate gene expression during oogenesis 2.1. Gal4/UAS Extensive Gal4/UAS tools are available for manipulating gene expression in germline and somatic cells of egg chambers. However, using the right UAS vector is critical. The original UASt vectors include P element ends for transformation and work very well in the somatic cells of the ovary; however, UASt-mediated expression is extremely poor in germline cells. Pernille Rørth [ 37 ] determined that the basal promoter from the hsp70 gene used in UASt is not expressed in germline cells. She produced new UASp vectors that instead use the basal promoter from the P element transposase gene. UASp vectors are included in the T. Murphy Collection of Gateway vectors at the Bloomington DGRC. A version of UASp that includes the attB sequence for phiC31-mediated integration was reported recently [ 38 ]. The UASt vector is available at the DGRC in Bloomington, and versions of UASt with the attB recombination sequence are available at FlyC31 . New UAS, attB vectors made in the Rubin lab at Janelia Farm are optimized for expression in the nervous system [ 39 ]; however, they work very well in follicle cells and also support expression in germline cells. Rubin lab vectors are listed on their website and are available at Addgene . A multitude of Gal4 lines is available from stock centers and labs that are useful for driving expression in specific cells of the ovary. The best source for comprehensive general information is the Gal4 page of the Bloomington Drosophila Stock Center (BDSC) , although this page has limited information on expression. Many of these lines are derived from Gal4 enhancer trap screens aimed at identifying lines expressed in during oogenesis and early embryogenesis [ 40 , 41 ]. Commonly used drivers for oogenesis research are listed in Table 1 , including lines for expression in the stem cell niche (germarium), germline cells and follicle cells. Gal4 expression from the nanos promoter is in germline stem cells, then recedes in young egg chambers and resumes later in oogenesis. The otu promoter produces more uniform expression in the germarium and egg chambers, but tapers off toward the end of oogenesis. A line called Maternal Triple Driver (MTD)-Gal4 has three Gal4 constructs ( P{otu-GAL4::VP16.R}1, w*; P{GAL4-nos.NGT}40; P{GAL4::VP16-nos.UTR}MVD1 ), providing robust germline expression throughout oogenesis [ 42 ]. For germline expression outside the germarium (starting in stage 2), the Matα-TubGal4 (MAT) line is ideal. Most of the lines driving expression in somatic follicle cells are also expressed in other tissues of the fly. An exception is Vm26a-Gal4, which has the promoter from the vitelline membrane 26Aa gene and is specific for follicle cells in stage 10–14 egg chambers [ 36 ]. A curious aspect of Gal4/UAS expression in egg chambers is frequent mosaicism. A group of follicle cells with one level of reporter expression is often adjacent to another group with a very different level. Nurse cells can also display different levels of expression [ 43 ]. The explanation for follicle cell mosaicism may involve epigenetic marks inherited by progeny during mitotic proliferation [ 44 ], though the level of variation in expression is somewhat masked by intercellular movement of proteins between follicle cells through ring canals [ 20 ]. The probability of mosaic expression levels should be taken into consideration when interpreting results of an experiment involving Gal4-mediated expression. A method to impose an additional layer of temporal control on Gal4 driven gene expression is the TARGET system [ 45 ]. TARGET relies on ubiquitous expression of Gal80ts, a specific, temperature sensitive repressor of Gal4, to inhibit Gal4 driven transgene expression at 18° C, the permissive temperature for Gal80ts. The Gal80ts-mediated repression is relieved by shifting flies to the restrictive temperature for Gal80ts (~29° C), which allows Gal4 to activate transcription. This method can works follicle cells [ 46 ], but in our experience Gal80ts is not able to repress strong Gal4 expression from Matα-TubGal4 in the germline. Repression of weaker germline Gal4 drivers may be possible; however, but this needs further testing. 2.2. Promoter fusions For rescue experiments or screening backgrounds, it can be desirable to have gene expression under the control of a promoter directly fused to a cDNA rather than using the bipartite Gal4/UAS system. Several vectors are available with promoters for germline-specific expression, or for ubiquitous expression in egg chambers ( Table 2 ). Vectors using the promoter from the α-Tubulin at 67C ( αTub67C ) gene direct strong, uniform expression in germline cells beginning in stage 2 egg chambers. The ovarian tumor ( otu ) promoter produces low to moderate levels of germline expression throughout oogenesis, and the hsp26 promoter produces moderate levels of expression starting in stage 8 egg chambers. The nanos ( nos ) or vasa ( vas ) promoter directs expression in primordial germ cells in developing animals, and in the germline cells of the adult germarium. Expression is low in early egg chambers, and high in late stages. For ubiquitous expression, the Ubiquitin-63E ( Ubi-p63E ) or spaghetti-squash promoter is a good choice. The α-Tubulin at 84B ( αTub84B ) promoter also provides expression in both germline and follicle cells, although expression is lower in the germline.

3.

Genetic methods

The greatest strength of Drosophila as an experimental system is the ability to use genetic analysis to understand cellular and developmental processes. Work from many labs during the past decades has resulted in remarkable innovations in the genetic tools available to study the genes and developmental mechanisms controlling oogenesis. We present here the basis for classical genetic approaches, and also summarize more recently developed techniques that rely on transgenic manipulations. 3.1. Classical genetics: female-sterile mutations Initial genetic approaches to study oogenesis focused on screens for recessive loss-of-function mutations affecting female fertility. The two broad classes of mutations affecting female fertility are female-sterile (fs) mutations and maternal-effect lethal mutations (mel). By definition, female-sterile mutations cause a defect during oogenesis such that normal eggs are not produced. In contrast, females bearing maternal-effect lethal mutations are able to produce normal eggs, indicating that oogenesis is able to proceed normally, but these eggs fail to produce viable offspring due to the lack of a gene product essential for embryonic development that is normally provided to the egg during oogenesis. Maternal-effect lethal mutations have been invaluable in understanding the mechanisms of early embryonic development. Screening for recessive female-sterile and/or maternal-effect lethal mutations requires an F 3 screen, where mutagenized chromosomes are balanced, made homozygous, and tested for effects on fertility. This task therefore requires substantial effort, and several large-scale of this type were conducted in 1970s and 1980s. As a result of these efforts, most genes that can be mutated to specifically affect female fertility have likely been identified [ 47 – 53 ]. 3.2.

Mosaic analysis

While female-sterile screens succeeded in isolating many genes that are critical for female fertility, the majority of genes that function in oogenesis are essential genes for which loss of function mutations do not produce a female-sterile phenotype. The study of these genes requires a means to inactivate or knock down gene function specifically in the ovary without compromising essential functions elsewhere in the fly. Historically, three experimental approaches have been developed to engineer genetically mosaic ovaries: pole cell transplantation, mitotic recombination, and tissue-specific RNAi. The latter two approaches are far easier to carry out and are more commonly used, but transplantation experiments can be useful in certain circumstances. 3.2.1. Pole cell transplantation Primordial germ cells form early in embryonic development and are the first embryonic cells to undergo cellularization. This allows them to be unambiguously identified in the embryo, and they can be transplanted from one embryo to another using micromanipulation. If the recipient embryo is derived from a strain lacking a functional germline (usually by carrying the dominant female-sterile gene ovo D1 ), then the resulting germline will be formed from the transplanted germ cells. Transplanted pole cells give rise to only germline tissue [ 54 , 55 ], so this technique can be used to determine whether a gene affecting oogenesis functions in the germ cells or the associated somatic cells. Experiments using such approaches led to important conclusions regarding sex determination and intercellular communication in Drosophila. A recent publication [ 56 ] describes an updated protocol on the technique that was originally developed in the 1970s [ 57 , 58 ]. 3.2.2. Mitotic recombination A versatile and widely used system to generate genetic mosaics in oogenesis is mitotic recombination. In this system, flies heterozygous for a mutation of interest are induced to undergo recombination during the G2 phase of the cell cycle so that inter-homolog exchange distal to a centromere will give rise to genetically distinct daughter cells. The wild-type chromosome typically includes a marker or reporter gene to allow for identification of recombined cells ( Figure 2 ). When mitotic recombination was first used to generate genetic mosaics, recombination was induced by exposing flies to ionizing radiation [ 59 ]. This method is relatively inefficient, however, and the introduction of the yeast FLP/FRT site-specific recombination system into flies vastly improved the efficiency of genetic mosaic analysis [ 60 ]. In this system, the yeast FLP enzyme acts on two F LP r ecombination t argets (FRTs) and catalyzes DNA strand exchange. Transgenes carrying FRT sequences have been recovered in centrosome-proximal positions on all five chromosome arms, enabling efficient mitotic recombination schemes to generate homozygous mutant clones for essentially all Drosophila genes [ 61 , 62 ]. Identifying mutant germline mosaics can be done genetically using Dominant Female Sterile (DFS) method or visually using reporters.

FLP-DFS system

The most efficient method to identify germline clones for later-staged egg chambers is the FLP-DFS technique [ 63 – 65 ]. This method uses a genetic selection to specifically mark germline mosaics resulting from mitotic recombination. The selection relies on the dominant female-sterile mutation ovo D1 , which when heterozygous results in an early arrest of germline development. In FLP-DFS, the experiment is designed so that ovo D1 and the mutation are on FRT-bearing chromosomes that are trans-heterozygous with each other. When recombination is induced in germ line stem cells or their progenitors, germ cells will be generated that are homozygous for the mutation of interest and, importantly, lack the ovo D1 mutation. The only egg chambers that survive to later stages are the non- ovo D1 egg chambers that are homozygous for the mutation of interest. Screens using elegant versions of this approach identified several genes involved in oocyte specification and development in [ 66 – 68 ].

Other marking systems

While the FLP-DFS system is an efficient means of generating late-stage germline clones, it is not useful for identifying germline clones during early oogenesis or for identifying clones in somatic cells. Instead, mosaics in these cell types can be identified using a variety of cytological markers. The simplest and most commonly used marking system for genetic mosaic analysis is negative marking, where mutant cells are marked by loss of a marker. Among these markers, nuclear-localized GFP expressed from the ubiquitin promoter is most commonly used, but other markers, including ubiquitously expressed β-galactosidase and the myc epitope fused to either nuclear or membrane localization domains can also be useful when GFP fluorescence is not desirable [ 62 ]. Chromosomes with these markers recombined onto the commonly used FRTs are available at the Bloomington stock center. It is important to note that the syncytial nature of ovarian cells can have important consequences for negative marking strategies. As mentioned in Section 1.1, follicle cells remain interconnected in syncytial nests of varying size, and small proteins can readily diffuse between connected follicle cells [ 20 ]. Thus it is probable that in when homozygous mutant cells are marked by loss of GFP, some GFP may diffuse from wild-type GFP producing cells to the neighboring mutant cells that lack the GFP transgene. Similarly, wild-type gene product may (or may not, depending on its ability to diffuse) move from a genetically wild-type cell to a homozygous mutant cell. The extent to which this is a concern in clonal analysis of follicle cells depends on the ability of the marker and the wild-type gene product to diffuse (for a more complete discussion of these issues, see [ 21 ]). In addition to negative marking, the MARCM system (Mosaic analysis with a repressible clonal marker) allows mutant cells to be positively marked by a UAS reporter transgene [ 69 ]. In MARCM, mitotic recombination is induced in a background where both Gal4 and its repressor Gal80 are ubiquitously expressed. Reporter gene expression is repressed by Gal80, which is expressed from a transgene that is heterozygous with the mutant of interest. Mitotic recombination results in progeny cells that are homozygous for the mutant and lack the Gal80 transgene; only these cells will express the UAS reporter. While this system is more complicated to set up, many stocks are available at Bloomington to facilitate MARCM crossing schemes ( MARCM stocks ). A further advantage of MARCM is that other UAS transgenes in addition to visible markers can be incorporated into MARCM experiments. This has been done in ovarian follicle cells for example, where mutant clones of a vacuolar ATPase were tested for their ability to suppress the effects of UAS-driven constructs that expressed activated forms of the Notch signaling protein [ 70 ]. In addition, it is possible to use biologically informative reporters as clonal markers; for example, a probe such as UAS-GFP::tubulin can be used as both a clonal marker and as a label for the microtubule cytoskeleton.

Lineage analysis

Mitotic recombination can also be used to trace cell lineage in a tissue as the cells divide. Lineage tracing experiments have been used to understand how somatic and germline stem cells contribute to ovarian tissue homeostasis (e.g., [ 71 , 72 ]. Several techniques based on FLP/FRT recombination have been developed that allow distinct labeling of mitotic lineages in oogenesis. The “X-15” lineage tracing system relies on mitotic recombination between FRT sequences to fuse the strong, ubiquitous α-tub84B promoter with the E. coli LacZ gene [ 73 ]. When recombination is induced during mitosis, one lineage inherits the promoter- LacZ fusion and will express high levels of β-galactosidase, which can be easily detected using either an X-gal staining reaction or immunofluorescence. Two additional lineage tracing systems, the twin spot generator (TSG; [ 74 ]) and twin spot MARCM [ 75 ] have been recently developed in which both cell lineages resulting from mitotic recombination are labeled. The twin spot generator is similar to the X-15 system in concept, but in this case FRT recombination reconstitutes ubiquitously driven RFP and GFP transgenes in the two lineages. Twin spot MARCM functions similarly to conventional MARCM, except that instead of Gal80-mediated repression, GFP and RFP are repressed by synthetic microRNAs in un-recombined cells. It is also possible to use the twin spot marking systems to compare wild type and mutant lineages, but only if a gene of interest is located distal to the FRT and transgene sites. Currently the twin spot MARCM transgenes are set up on chromosome arm 2L, while twin-spot generator transgenes are available at centromere-proximal locations on chromosome arms 2L, 3L, and 3R [ 74 ]. 3.2.3. RNA interference RNAi in Drosophila has become an immensely powerful tool, and at least three genomic-scale projects have been undertaken with the goal of establishing transgenic RNAi lines for essentially all Drosophila genes [ 76 , 77 ] and RNAi FLY , unpublished (available lines are listed at the following sites: ( VDRC , TRiP , and RNAi FLY ). For many cell types in Drosophila, RNAi is efficiently induced by expressing an inverted-repeat “hairpin” RNA [ 78 ]. The Vienna and RNAi FLY transgenic RNAi collection consists of transgenes that express hairpin RNAs [ 76 ] under UAS control. However, hairpin RNAs do not effectively silence genes in the female germline, limiting their usefulness in studying oogenesis. More recently, a novel transgenic RNAi system has been developed that allows potent gene silencing in all cell types, including ovarian germ cells [ 79 ]. This method was designed to express short RNA hairpins in a microRNA backbone; these RNAs appear to be more efficient in gene knockdown [ 80 ]. Experimental tests of this system demonstrated that the short micro-RNAs (shRNAs) produced a potent knockdown of target genes in all cell types examined, including ovarian germ cells [ 79 ]. A new large-scale transgenic RNAi collection is being generated using the shRNA strategy, and these lines are also available at TRiP . Effective RNAi silencing in the germline enabled a large-scale screen for genes involved in ovarian stem cell regulation [ 81 ]. By testing RNAi lines driven by either MTD-Gal4, which drives UAS-RNAi expression beginning in stem cells and their progenitors, or Matα-TubGal4, which begins driving expression after germline cyst formation is complete, it was possible to identify genes that function in the germarium, outside the germarium, or both.