embryo transformation

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samar El Kholy
samar El Kholy's picture
embryo transformation

hallo,
i have problems in injecting drosophila embryos . i use double -sided tap (non toxic) and dechorionate embryos manully but embryos dried even after 5 min. andwhen i put oil embryos becime not fix and very difficilt to inject .

Any suggession?

sezzat

Marina Fomin
Marina Fomin's picture
What if you put PBS or Saline

What if you put PBS or Saline Sol. on the embryo?
It is just a suggestion, I don't have expirience with injection of drosophila emb. Only chiken or mouse.

Tony Rook
Tony Rook's picture
sezzat:

sezzat:

Here are a few references that may help...

Characterization of fluidic microassembly for immobilization and positioning of Drosophila embryos in 2-D arrays Sensors and Actuators A: Physical Volume 114, Issues 2-3, 1 September 2004, Pages 191-196
doi:10.1016/j.sna.2003.11.021

Abstract:
A technique for the positioning and immobilization of Drosophila embryos in 2-D arrays for use in high throughput microinjection experiments has been characterized. The method is based on fluidic microassembly, and immobilization yield, the number of misplaced embryos, alignment properties, and adhesion force of the embryos have been measured for samples with four different pad geometries. For samples with 250 μm×400 μm sized rectangular pads, an immobilization yield of 85% was achieved. The adhesion force of the embryos was estimated at 36±22 μN. A substantial amount of clustering was, however, observed. By reducing the pad size to 200 μm×200 μm and changing the pad pitch and shape, the number of misplaced embryos was reduced to less than 5%. The adhesion force of embryos immobilized at the smaller pads was estimated at 14±5.5 μN, resulting in lower immobilization yield for these samples. The self-assembly process facilitates rotational alignment to some degree, and an average of 40% of the immobilized embryos align within ±9° of the symmetry axis of the immobilization sites.

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Bernstein R.W., Xiaojing Zhang Zappe S., Fish M., Scott M., Solgaard O. Characterization of Drosophila embryos immobilized by fluidic microassembly. TRANSDUCERS, Solid-State Sensors, Actuators and Microsystems, 12th International Conference on, 2003
Publication Date: 8-12 June 2003
Volume: 2, On page(s): 987- 990 vol.2
ISBN: 0-7803-7731-1

Abstract:
A technique for positioning and immobilization of Drosophila embryos in 2-D arrays for use in high throughput microinjection experiments has been characterized. The method is based on fluidic microassembly and immobilization yield, the number of misplaced embryos, and adhesion force of the embryos have been measured for samples with two different pad geomtries. For samples with 250 /spl mu/m/spl times/400 /spl mu/m sized pads an immobilization yield of 85% was achieved. The adhesion force of the embryos was estimated at 36 /spl mu/N/spl plusmn/22 /spl mu/N. A substantial amount of clustering was, however, observed. By reducing the pad size to 200 /spl mu/m/spl times/200 /spl mu/m and changing the pad pitch the number of misplaced embryos was reduced to less then 5%. The immobilization yield was, however, lower for these samples. The adhesion force of embryos immobilized at the smaller pads was estimated at 14 /spl mu/N/spl plusmn/5.5 /spl mu/N.

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Alain Debec, Robert F. Kalpin, Douglas R. Daily, Patrick D. McCallum, Wendy F. Rothwell, and William Sullivan. Live Analysis of Free Centrosomes in Normal and Aphidicolin-treated Drosophila Embryos. The Journal of Cell Biology, Volume 134, Number 1, July 1996 103-115.

Abstract:
In a number of embryonic systems, centrosomes that have lost their association with the nuclear envelope and spindle maintain their ability to duplicate and induce astral microtubules. To identify additional activities of free centrosomes, we monitored
astral microtubule dynamics by injecting living syncytial Drosophila embryos with fluorescently labeled tubulin. Our recordings follow multiple rounds of free centrosome duplication and separation during the cortical divisions. The rate and distance of free sister centrosome separation corresponds well with the initial phase of associated centrosome separation. However, the later phase of separation observed for centrosomes associated with a spindle (anaphase B) does not occur. Free centrosome separation regularly occurs on a plane
parallel to the plasma membrane. While previous work
demonstrated that centrosomes influence cytoskeletal
dynamics, this observation suggests that the cortical cytoskeleton regulates the orientation of centrosome separation.
Although free centrosomes do not form spindles, they display relatively normal cell cycle-dependent modulations of their astral microtubules. In addition, free centrosome duplication, separation, and modulation of microtubule dynamics often occur in synchrony with neighboring associated centrosomes. These observations suggest that free centrosomes respond normally
to local nuclear division signals. Disruption of the cortical
nuclear divisions with aphidicolin supports this conclusion;
large numbers of abnormal nuclei recede into the interior while their centrosomes remain on the cortex. Following individual free centrosomes through multiple focal planes for 45 min after the injection of aphidicolin reveals that they do not undergo normal
modulation of their astral dynamics nor do they undergo
multiple rounds of duplication and separation. We conclude that in the absence of normally dividing cortical nuclei many centrosome activities are disrupted and centrosome duplication is extensively delayed. This indicates the presence of a feedback mechanism that creates a dependency relationship between the cortical nuclear cycles and the centrosome cycles.

Paraphrased Method from this reference -

"The in vivo analysis of nuclear and centrosome behavior was
accomplished by microinjecting fluorescently labeled histones
and tubulin into embryos during the syncytial cortical divisions
(Kellogg et al., 1988; Minden et al., 1989). The embryos were
prepared for microinjection by hand dechorionation and
mounting on a coverslip with a thin film of glue (Minden et al.,
1989). Observations and time-lapse recordings were made on
an Olympus IMT2 microscope equipped with a Bio-Rad MRC
600 confocal imaging system.

A 100-txg/ml solution of aphidicolin dissolved in a 0.5% DMSO,
5 mM KCI, 0.1 mM sodium phosphate (pH 6.8) solution was
used to inhibit DNA synthesis. 1-h collections of embryos aged
for 30 min were injected with either rhodamine-labeled
histones or rhodamine-labeled tubulin.

Once the nuclei migrated to the cortex, these embryos were
injected with 100 ixg/rrd aphidicolin. The microtubule and
nuclear dynamics in these embryos were observed for up to
1 h after the injection."

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References cited in the above reference Debet at al. 1996:

Kellogg, D.R., T.J. Mitchison, and B.M. Alberts. 1988. Behaviour of microtubules and actin filaments in living Drosophila embryos. Development, Vol 103, Issue 4 675-686.

Abstract:
We describe the preparation of novel fluorescent derivatives of rabbit muscle actin and bovine tubulin, and the use of these derivatives to study the behaviour of actin filaments and microtubules in living Drosophila embryos, in which the nuclei divide at intervals of 8 to 21 min. The fluorescently labelled proteins appear to function normally in vitro and in vivo, and they allow continuous observation of the cytoskeleton in living embryos without perturbing development. By coinjecting labelled actin and tubulin into the early syncytial embryo, the spatial relationships between the distinct filament networks that they form can be followed second by second. The dynamic rearrangements of actin filaments and microtubules observed confirms and extends results obtained from previous studies, in which fixation techniques and specific staining were used to visualize the cytoskeleton in the Drosophila embryo. However, no tested fixation method produces an exact representation of the in vivo microtubule distribution.

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Minden, J.S., D.A. Agard, J.W. Sedat, and B.M. Alberts. 1989. Direct cell lineage analysis in Drosophila melanogaster by time-lapse, three-dimensional optical microscopy of living embryos. J. Cell Biol. 109:505-516.

Abstract:
One of the first signs of cell differentiation in the Drosophila melanogaster embryo occurs 3 h after fertilization, when discrete groups of cells enter their fourteenth mitosis in a spatially and temporally patterned manner creating mitotic domains (Foe, V. E. and G. M. Odell, 1989, Am. Zool. 29:617-652). To determine whether cell residency in a mitotic domain is determined solely by cell position in this early embryo, or whether cell lineage also has a role, we have developed a technique for directly analyzing the behavior of nuclei in living embryos. By microinjecting fluorescently labeled histones into the syncytial embryo, the movements and divisions of each nucleus were recorded without perturbing development by using a microscope equipped with a high resolution, charge-coupled device. Two types of developmental maps were generated from three-dimensional time-lapse recordings: one traced the lineage history of each nucleus from nuclear cycle 11 through nuclear cycle 14 in a small region of the embryo; the other recorded nuclear fate according to the timing and pattern of the 14th nuclear division. By comparing these lineage and fate maps for two embryos, we conclude that, at least for the examined area, the pattern of mitotic domain formation in Drosophila is determined by the position of each cell, with no effect of cell lineage.

________________________________________________________

Good luck!

Tony Rook
Tony Rook's picture
Here's another reference

Here's another reference which should be helpful -

SANTAMARIA, P. (1986). Injecting Eggs. In
Drosophila: A Practical Approach (ed. D. Roberts) , pp. 159-174.
Washington, DC: IRL Press.

Tony Rook
Tony Rook's picture
By the way sezzat...

By the way sezzat...

I found most of these references by using the Solutions Search tool on this site.

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Tony Rook
Tony Rook's picture
Just one more... in case you

Just one more... in case you want to get fancy and automate the process!

Wenhui Wang, Xinyu Liu, Danielle Gelinas, Brian Ciruna, Yu Sun. A Fully Automated Robotic System for Microinjection of Zebrafish Embryos. PLoS ONE 2(9): e862. doi:10.1371/journal.pone.0000862

Open Access Abstract:
Molecule screening at the single cell level, which is critical in molecular biology and drug discovery, requires that target molecules be introduced into single cells to permit cellular-function-targeted molecules to directly regulate cell development and their functions to be quantified. Several technologies exist for introducing foreign materials into a cell, such as electroporation [1], viral vectors [2], gene gun [3], ultrasonics [4], and MEMS-based injection [5][6]. Compared to these techniques, microinjection with a single glass micropipette remains the most effective in terms of cell damage, cell viability, cell waste, effectiveness of delivering macromolecules, specificity, and freedom from concerns about phenotype alteration. However, in order to enable fast, precise, and robust screening for molecular targets, the state-of-the-art manual injection must be replaced with fully automated operation.

For testing cellular responses to molecular targets and to obtain statistically significant data, the injection of thousands of cells needs to be conducted within a short time window (e.g., within 1.5 hr after fertilization, before the 16-cell stage for zebrafish embryo injection). Manual injection is not only slow; the laborious task of manual injection easily causes fatigue in injection technicians and hinders performance consistency and success rates. Efforts in automating cell injection have been continuous, resulting in a visually servoed system [7], a semi-automated system [8], and many tele-operated systems [9][13], to name just a few. These systems are limited in throughput and reproducibility as operator input (e.g., locating features and destinations) or operator involvement (e.g., switching from one cell to another or injector alignment) is still required.

Among many biological models, the zebrafish has emerged as an important model organism for developmental genetic studies as well as for drug discovery [14][15]. Zebrafish embryonic development is remarkably similar to that of humans; however, zebrafish embryos are laid and fertilized externally, they develop rapidly, and the embryos are transparent (Figure 1), making it convenient to observe the movement and fate of individual cells during embryonic development [16]. Molecular and genetic analyses of zebrafish embryogenesis depend on the injection of foreign materials into early zebrafish embryos [17]. DNA injection is used to generate transgenic zebrafish lines, mRNA injection is used to overexpress gene-products in zebrafish embryos, and reverse genetic or loss-of-gene-function studies require the injection of antisense morpholino-modified oligonucleotides (morpholinos or MOs) to specifically inhibit RNA splicing and/or translation in vivo [18].

Despite their relatively large size (~600 m or ~1.2 mm including chorion), zebrafish embryos have a delicate structure and can be easily damaged, making automated, high-throughput injection difficult. Specific challenges include: (i) to quickly (i.e., in seconds) immobilize a large number of zebrafish embryos; (ii) to automatically, robustly identify cell structures for vision-based position control and account for size differences across embryos; and (iii) to coordinately control two microrobots to maximize operation speed. Addressing these challenges, the objective of this research was thus to develop an effective massive sample preparation method and create a system that is capable of injecting a large number of embryos in the short time window. In this paper, a microrobotic system for zebrafish embryo injection is presented, featuring full automation, high-speed sample immobilization, and high survival, success, and phenotypic rates.

samar El Kholy
samar El Kholy's picture
Marina Fomin wrote:What if

Marina Fomin wrote:

What if you put PBS or Saline Sol. on the embryo?
It is just a suggestion, I don't have expirience with injection of drosophila emb. Only chiken or mouse.

Do you see that using holding pipette as used in mouse injection will help me to immobilize Drosophila embryos because once i put oil embryos moved and the stick tap become without effect?
Thanks a lot

Marina Fomin
Marina Fomin's picture
I guess so... Holding pipette

I guess so... Holding pipette can help you a lot if your have a manipulator for injections. You can make it any size you want. Mouse emb. we inject in drop of medium covered with oil. Again, I don't have personal experience with Drosophila emb.