Abstract
Transgenic models are invaluable tools for researching retinal degenerative disease mechanisms. However, they are time-consuming and expensive to generate and maintain. We have developed an alternative to transgenic rodent models of retinal degeneration using transgenic Xenopus laevis. We have optimized this system to allow rapid analysis of transgene effects in primary transgenic animals, thereby providing an alternative to establishing transgenic lines, and simultaneously allowing rigorous comparisons between the effects of different transgenes.
Key words
1 Introduction
We have developed techniques for generating and analyzing transgenic Xenopus laevis models of retinal degeneration (RD) based on mutations in the rhodopsin gene responsible for retinitis pigmentosa in humans (1–9); we are also currently applying these techniques to the analysis of other transgenes in our laboratory. In addition, we (and others) have applied these and related techniques to studies of protein localization in photoreceptors (5–19), and to the analysis of promoter activities (20–27). In general, these techniques differ from those typically used for analysis of transgenic rodents in that founder (“F0” or “primary”) transgenic animals are analyzed without further breeding to create transgenic lines, and the effects of multiple transgenes can be compared in a single experiment. Transgenic X. laevis are generated by injection of transgenic sperm into unfertilized eggs (the REMI/nuclear transplantation method) (28), followed by an antibiotic selection protocol to eliminate non-transgenic embryos (29). For antibiotic selection purposes, the DNA construct must contain a second transgene—a neomycin resistance cassette based on the aphA-2 gene capable of functioning in eukaryotic cells, such as that found in the eGFP-N1 vector (Clontech). The primary transgenic animals generated by these methods are non-chimeric and can therefore be analyzed without further breeding (28). In our experiments, the animals are typically sacrificed and analyzed two weeks after a set of injections, thereby allowing an entire experiment to be completed in under a month.
Using these techniques, two experienced individuals working together can easily generate more than 200 transgenic animals in a single day. A typical experiment would involve at least two transgenes—for example, a comparison of the effects of a wild-type rhodopsin cDNA and a rhodopsin mutant. However, we have often compared multiple transgenes (up to eight) (6). The use of a wild-type rhodopsin cDNA allows us to control for the effects of rhodopsin overexpression, although in contrast to results obtained in mice (30, 31), these effects are typically quite minimal.
In addition, transgenic X. laevis containing two transgenes are also readily produced by simultaneously injecting two different transgene constructs (3)—the high success rate is likely due to high transgene copy numbers obtained using this method.
Because each resulting transgenic animal contains unique transgene integration sites and copy numbers, expression levels vary considerably between animals (typically over a range of two orders of magnitude for wild-type rhodopsin), with the highest expression levels approaching or exceeding 50% of total rhodopsin being least common (5–7). Given a sufficient yield of primary transgenic animals, this allows investigators to survey transgene effects at a variety of expression levels, and also to monitor the effects of mutations on protein expression levels (5–8). This requires a relatively high “N,” but this is generally easily achieved.
However, this non-normal distribution of expression levels, combined with the likelihood that a threshold expression level is required to achieve a phenotype, requires that special considerations must be made for the analysis of transgene effects and RD—in particular, nonparametric statistical methods (which do not assume normally distributed data) should be employed when analyzing primary transgenic animals (5–8).
One confounding aspect of the system is the presence of position effects that result in transgene silencing in a relatively high proportion of transgenic animals. This silencing can be transient (on and off) or relatively permanent, and can affect a subset or virtually all cells in an animal (15). We have found that position effects can be minimized by two techniques that can be applied in combination: the use of antibiotic selection (29) (presumably because the neomycin resistance cassette is subject to similar silencing and therefore embryos with strong position effects do not survive), and the use of a chicken β-globin “double insulator” sequence in transgene constructs (32, 33).
Although we have attempted to duplicate these procedures in the diploid species Xenopus tropicalis, the resulting yield of primary transgenic animals is prohibitively low in our hands. This is likely due to the relatively small size of X. tropicalis eggs, which sustain comparatively greater damage on injection. We have also attempted to duplicate these procedures using other published methods for generation of transgenic X. laevis, including ϕC31 integrase (34) and ISce1-mediated transgene integration (35, 36) methods. In our hands, these generate high yields of primary transgenic animals consistent with the original reports. However, the expression levels are insufficient to allow for the development of an RD phenotype in a significant proportion of the animals.
The procedures we have described can also be used to generate transgenic lines with consistent RD phenotypes (2, 37). Our standard approach is to initially conduct experiments with primary transgenic animals while setting aside a population to rear to sexual maturity (requiring 8 months to 1 year).
Here we present protocols for generation of transgenic embryos by nuclear transplantation, antibiotic selection of transgenic embryos, and dot blot analysis of total rod opsin levels to monitor RD. Other valuable resources for X. laevis transgenesis procedures are also available (38, 39).
2 Materials
2.1 Generation of Transgenic X. laevis Embryos
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1.
Special equipment and materials required: Two high-quality dissecting microscopes (e.g., Zeiss Stemi), two micromanipulators (e.g., Narishige MM-3 or similar), a syringe pump (e.g., Harvard pump 11 plus) equipped with two 50 μl syringes, Tygon tubing, microcap pipettes, pipette puller (e.g., Flaming/Brown type pullers manufactured by Sutter Instruments), cold room, adult X. laevis females (Fig. 1a shows a typical injection setup).
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2.
Needles for microinjection—we use microcap pipettes (Drummond, 30 μl) using a single pull to achieve a taper length of approximately 1 cm, which we then break with tweezers to give a hypodermic-like profile, and an opening diameter of 60–80 μm (Fig. 1b, c).
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3.
Sperm nuclei—prepared according to Murray (40) as modified by Kroll and Amaya (28) and subsequently made up to a final concentration of 30% glycerol and 100 nuclei/nl, then aliquoted, and frozen at −80° for indefinite storage. We will typically do a prep using testes from 1 to 2 animals that will be sufficient for a large number of experiments.
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4.
High-Speed Egg Extract—prepared according to Murray (40) as modified by Kroll and Amaya (28), aliquoted, and frozen at −80°.
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5.
Human chorionic gonadotropin (Sigma Aldrich)—made up at 1 unit/μl.
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6.
Linearized transgene DNA—This can be prepared by either linearization of plasmids containing the transgenes with a restriction enzyme followed by gel purification (we use kits available from Qiagen) or amplification of the necessary sequence by long accurate PCR followed by similar gel purification. The final prep should be diluted to 75 ng/μl in dH2O. If the antibiotic selection protocol is to be used, it is critical that a neomycin-resistance cassette is present in the transgene construct, and is not disrupted by the plasmid linearization process. It is also critical to eliminate all circular DNA, and to use DNA at the specified concentration (see Note 1).
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7.
Mark’s modified ringer (MMR): 100 mM NaCl, 2 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 5 mM HEPES pH 7.4 and autoclaved. We prepare 1 and 20× stocks.
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8.
Tadpole Ringer: 10 mM NaCl, 0.2 mM KCl, 0.1 mM MgCl2, 0.2 mM CaCl2. We prepare as 100× stock.
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9.
2% Cysteine solution: 1 g of cysteine hydrochloride added to 50 ml of 1× MMR, pH adjusted to 8.0 with 2N NaOH, freshly prepared on the day needed.
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10.
Sperm dilution buffer (SDB): 250 mM sucrose, 75 mM KCl, 0.5 mM spermidine trihydrochloride, 0.2 mM spermine tetrahydrochloride; titrated to pH 7.4 with NaOH.
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11.
Ficoll solutions: 0.1× MMR + 6% Ficoll 400, and 0.4× MMR + 6% Ficoll 400, filter sterilized.
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Gentamycin stock solution: 10 mg/ml gentamycin.
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Agarose injection plates: 2% agarose dissolved in dH2O and autoclaved. Pour into 50 mm petri dishes and float 2 × 2 cm pieces of autoclaved silicone (cut from a silicone baking sheet or a similar product) in the center of the dish. Once the agarose has set, cover the surface with sterile 1× MMR, replace lids, and seal with parafilm for storage at 4°C.
2.2 Antibiotic Selection of Transgenic Embryos
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1.
Translucent or transparent flat-bottomed plastic bins suitable for rearing tadpoles, such as Nasco Flex-tanks. Glass tanks can also be used.
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2.
G418 (Geneticin) (Gibco-BRL).
2.3 Blot Analysis of Retinal Degeneration
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Dot blot apparatus (for example Biorad Bio-Dot).
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2.
LI-COR Odyssey imaging system.
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3.
Nylon membrane (e.g., Immobillon-P, Millipore).
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4.
Miniature homogenizer (e.g., Kontes Pellet Pestle).
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Mini-scalpel and forceps or similar dissection tools (e.g., Beaver Microsharp or a similar one).
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6.
Tricaine (MS-222) anesthetic.
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7.
Modified SDS-PAGE loading buffer (standard SDS-PAGE loading buffer diluted 2× with phosphate-buffered saline (PBS), and containing 1 mM PMSF and 2 mM EDTA). For 1 ml, combine 0.5 ml of PBS, 0.5 ml of SDS-PAGE loading buffer (5% SDS, 0.1 M Tris HCl pH 6.8, 40% sucrose, and 0.1% Bromophenol blue), 40 μl of 2-mercaptoethanol, 4 μl of 0.5 M EDTA pH 8.0, and 10 μl of 100 mM PMSF.
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8.
20 mM Sodium Phosphate pH 7.5.
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9.
Hybridoma B630N cell culture supernatant (a monoclonal anti-rod opsin antibody with broad cross-reactivity available from Dr. W. Clay Smith, University of Florida).
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10.
IR-Dye 800 conjugated goat anti-mouse secondary antibody (LICOR).
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11.
PBS, powdered milk, Tween-20, and SDS.
3 Methods
3.1 Generation of Transgenic X. laevis Embryos
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1.
Prepare linearized transgene DNA.
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2.
Two days prior to experiment: late in the day, inject three or more female X. laevis with 50 units of human chorionic gonadotropin (subcutaneous injection). The females are subsequently kept in 18°C dH2O containing 10 mM NaCl.
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3.
One day prior to experiment: Late in the day, reinject the female X. laevis with 700 units of human chorionic gonadotropin.
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4.
On the day of the experiment—Prepare fresh cysteine solution, and dilute the restriction enzyme (the same enzyme used for linearization of the transgene construct) to 0.1 unit/μl with SDB. Thaw aliquots of egg extract and sperm nuclei and store with diluted restriction enzyme on ice (see Note 2).
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5.
Ensure that the female X. laevis are laying eggs, and transfer them to clean tanks.
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6.
Steps 7–14 are conveniently split between two researchers. One researcher can carry out steps 7–9 while the second carries out steps 10–14. An experienced individual can perform all steps simultaneously. However, the goal is for steps 9 and 14 to be completed simultaneously.
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7.
Squeeze eggs into a 10 cm petri dish containing 1× MMR. Squeezing frogs correctly requires some instruction and practice, and should not cause any injury to the frogs. Do not use eggs that have been previously laid into the tank water. Try to obtain on the order of 800–1,600 eggs.
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8.
Replace the 1× MMR solution covering the eggs with 2% cysteine solution, and agitate the dish until the jelly coating is removed. Wash the eggs repeatedly in 1× MMR, removing any abnormal or lysing eggs in the process (at least three washes).
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9.
Remove the 1× MMR solution and silicone square from a sufficient number of agarose dishes (typically 2–4) and replace with 0.4× MMR + 6% Ficoll solution. Transfer the de-jellied eggs into the wells formed by the silicone squares, carefully forming a single layer of tightly packed eggs, while transferring as little solution as possible.
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10.
Add 2.5 μl of the linearized DNA constructs to two Eppendorf tubes (if two different constructs will be injected by the two researchers) (Fig. 1a).
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11.
Add 2 μl of sperm nuclei to each tube, and incubate for 5 min at 18°C.
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12.
Add 0.5 μl of diluted restriction enzyme, 3.5 μl egg extract, and 10 μl of SDB, mix gently, and incubate for 10 min at 18°C.
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13.
Dilute the reaction in SDB to give a final concentration of 0.3 nuclei/nl (see Note 3). Back-load two injection needles with the reactions, connect the needles to the syringe pump, and mount them in the micromanipulators. The injection needles should be mounted at an angle of 45° from horizontal. Set the syringe pump to deliver 36 μl/h (10 nl/s) (see Note 4).
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14.
Inject the eggs as quickly as possible, making sure to leave the needle in each egg for a total of 1 s. Eggs should be injected half way between the pole and equator, such that the needle enters perpendicularly, doing as little damage as possible (Fig. 1d–e). An experienced injector can inject at a rate of 40 eggs/minute, so two individuals can inject 1,600 eggs in approximately 20 min. After this point, both the eggs and reactions will begin to deteriorate in quality.
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15.
After the first set of injections are complete, set the plates aside, and resume the procedure at step 6 to perform another set of injections (if desired).
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16.
Monitor the injected eggs and watch for the start of cell division—this should be reasonably synchronized across the plate. At the second cell division, use a fire-polished Pasteur pipette to remove all the correctly dividing eggs (four cells) that can be identified (Fig. 1f), and place them in a dish of 0.1× MMR + 6% Ficoll containing 50 μg/ml gentamycin antibiotic. Under optimal conditions, 1/3 of the injected eggs will divide correctly, 1/3 will not divide, and 1/3 will divide abnormally due to receiving more than one sperm nucleus. Assess the numbers in each category, and adjust the dilution factor or quantity of nuclei added in further reactions accordingly.
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17.
After 24 h, transfer the surviving embryos to tadpole ringer for rearing (Fig. 1g), or proceed with the antibiotic selection protocol below if desired. Under optimal conditions, the majority of the correctly dividing embryos will survive the first 24 h, and the yield after 14 days will be reduced by approximately 1/3—the attrition is likely due to genomic damage. Approximately 1/3 of the embryos will be transgenic.
3.2 Antibiotic Selection of Transgenic Embryos
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1.
On the day following fertilization (gastrula or neurula stage), transfer the surviving embryos to 0.1× MMR containing 20 μg/ml G418. Place no more than 200 embryos per liter of antibiotic solution in a suitable container such as a Nasco Flex Tank, and maintain the tanks at 18°C.
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2.
Monitor the embryos, continuously removing any dead embryos—after 120 h, a significant proportion of the embryos will be developmentally delayed, arrested, or dead. For the purposes of troubleshooting it may be useful to divide initial experiments into selected and nonselected groups so that the effects of the G418 treatment can be differentiated from those due to normal attrition. Occasionally shorter treatments (96 h) or longer treatments (up to 144 h) are necessary before selection effects are apparent.
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3.
When healthy and developmentally delayed embryos can be clearly differentiated (Fig. 1h, i), the healthy embryos can be separated manually (regardless of whether the selection has progressed to completion), transferred to 0.1× MMR or 1× tadpole ringer, and allowed to develop to the stage required for analysis (e.g., stage 49–50, 14 days following fertilization). At this point, the animals are transferred to normal housing conditions (clear plastic tanks containing tadpole ringer, 18°C, with daily feeding of powdered frog chow). Typically, 20% of the embryos subjected to G418 selection will survive, and >90% of these will be transgenic.
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4.
Although this protocol usually results in >90% transgenic animals, it is still critical to confirm that the majority of animals are transgenic. This can be done by a variety of methods, the simplest being to screen the animals visually for fluorescence if the transgene encodes a fluorescent product such as a GFP fusion protein. Other suitable methods include PCR of genomic DNA, southern blot (15), co-injection of a transgene encoding a fluorescent protein (3), immuno-labeling of frozen sections from the contralateral eye with an antibody specific for the transgene product (5–7), or a dot- or western blot assay similar to that described below using an antibody specific for the transgene product (5–7) (we have used antibodies 1D4 and 2B2 for labeling our rhodopsin transgene products, as these antibodies do not label the endogenous X. laevis rhodopsin; we have also used anti-HA for HA-tagged transgene products (3)).
3.3 Dot Blot Analysis of Retinal Degeneration
The following procedures are optimized for a time point of 14 days post fertilization.
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Sacrifice the transgenic tadpoles by an overdose of Tricaine anesthetic (0.1% in 0.1× MMR).
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Enucleate one eye from each animal to be analyzed and place in an Eppendorf tube containing 100 μl of modified SDS-PAGE loading buffer. (The remaining eye can be used for other assays such as histology or RNA isolation. For histology, fix overnight in 4% paraformaldehyde in 0.1 M sodium phosphate buffer pH 7.5.)
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3.
Homogenize for 30 s, rinsing the homogenizer in dH2O between samples.
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4.
Once all samples are homogenized, centrifuge for 5 min at maximum speed in a benchtop microfuge (see Note 5).
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5.
Dilute each sample 300× using 20 mM sodium phosphate pH 7.5 (use a 96-well plate to aid organization) (see Note 3 and Note 6).
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6.
Load the dot blot apparatus with a sheet of membrane prepared according to the manufacturer’s instructions, and add 270 μl of sodium phosphate buffer per well (see Note 7).
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7.
Add 30 μl of each sample prepared in step 6 to each well prepared in step 7 (i.e., each sample is further diluted on loading into the apparatus, for a combined dilution of 3,000×) (see Note 3 and Note 6).
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8.
Draw the samples through the membrane using vacuum, followed by several rinses of dH2O.
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9.
Remove the membrane from the apparatus, rinse with dH2O, and air-dry.
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10.
Prepare the membrane for immunoblotting according to the manufacturer’s instructions (see Note 7).
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11.
Block with PBS containing 1% powdered milk for 30 min.
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12.
After rinsing with PBS/0.5% Tween, label overnight with B630N cell culture supernatant diluted 20× in PBS/0.5% Tween/0.1% powdered milk (see Note 3 and Note 6).
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13.
After rinsing with PBS/0.5% Tween, label with IRDye800 secondary antibody diluted 10,000× in PBS/0.1% powdered milk/0.5% Tween/0.02%SDS for 2–4 h. This antibody must be protected from all light exposure (see Note 3 and Note 6).
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14.
Image the dot blot using a LI-COR Odyssey imaging system. Quantify the signal from each sample—these measurements can be used for comparative purposes between animals and between groups of animals. In general, a low B630N signal (i.e., low rod opsin levels) is likely to be a result of missing or abnormal rod photoreceptors, and indicates RD—however this should be confirmed by a second assay (for example, histology performed on contralateral eyes) (Fig. 2). Alternatives to the Odyssey imaging system (such as chemiluminescence-based detection) could be employed. The Odyssey imaging system gives a broad linear response and is ideal for this purpose. However, nonparametric statistical methods (necessitated by the non-normal distribution of the resulting data; see introduction) require only ordinal data, and therefore a completely linear response is not critical, although every effort should be made to ensure that the assay response is as linear as possible. The assay can be easily optimized using serial-diluted samples obtained from non-transgenic eyes.
4 Notes
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1.
For troubleshooting purposes, it may be useful to use a GFP transgene with expression driven in all tissues by a promoter such as CMV.
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2.
From this point onward, it is advisable (but not critical) to carry out further steps in an 18°C cold room.
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3.
Dilutions and concentrations may need to be optimized.
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4.
Theoretically, it is optimal to inject an average of one sperm nucleus per egg, whereas we typically inject three. This number was arrived at empirically, and may reflect a tendency for sperm nuclei to leak from the injection site, and for a portion of the sperm nuclei to be nonviable.
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5.
These samples are also suitable for analysis by standard western blot.
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6.
The concentrations given result in the majority of samples lying within a linear response range of the assay in our hands.
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7.
For Immobilon-P, wet with methanol, and then rinse with dH2O.
References
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This research was funded by the Canadian Institutes for Health Research, and the Foundation Fighting Blindness (Canada).
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Tam, B.M., Lai, C.CL., Zong, Z., Moritz, O.L. (2012). Generation of Transgenic X. laevis Models of Retinal Degeneration. In: Weber, B., LANGMANN, T. (eds) Retinal Degeneration. Methods in Molecular Biology, vol 935. Humana Press, Totowa, NJ. https://doi.org/10.1007/978-1-62703-080-9_8
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DOI: https://doi.org/10.1007/978-1-62703-080-9_8
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