Information

Measuring transgenic gene expression with a microplate reader


I have been working on genetically engineering an E. coli strain to autotrophically oxidize arsenite into arsenate for bioremediation of arsenic contamination in groundwater. For my research, I have been finding ways to quantitatively measure how much arsenite the bacteria oxidizes over time. After researching ways to quantitatively measure arsenite being oxidized, I have concluded to use the method of Johnson and Pilson (https://www.sciencedirect.com/science/article/pii/S0003267072800059?via%3Dihub).

The arsenate that is formed from arsenite oxidation forms a molybdenum-blue complex with phosphate in the water. As the concentration of arsenate increases, the absorbance value of the water should increase proportionally.

Here was my plan: First, after growing my transgenic bacteria on an amp-plate, I would pipette out various concentrations of cells and arsenite in wells of a 96-well spot plate. I currently plan to use the Fisher Scientific Multiskan FC microplate photometer. I planned on incubating the cells while shaking at 37 C. The main problem I have come across is that my experiment would not be controlled. Over time, the growth of cells will change the absorbance value, not just the molybdenum-blue complex. I need to isolate the variable of just arsenite oxidation.

I have thought of a few possible solutions, but I feel as if I may be missing something crucial. When learning about microplate readers, I know that researchers can use them to measure gene expression. How do these researchers eliminate the variable of cell growth effects on absorbance values?

For one, I have thought of finding a way to filter the cells to one side of the well where they will not affect the absorbance values. Would it be possible for me to cut out a dialysis bag to where it is dipping into the well? In that case, the cells could be suspended in the well, oxidizing arsenite (if my hypothesis is correct).

Anyway, I really want to find a simple method for quantitatively measuring arsenite concentrations over time using a microplate reader without having cell growth affecting absorbance values.


Spectophotometry

Does the photometer allow using different wavelengths of light? Perhaps using a spectrophotometer might give you a control and your desired value, i.e. blue light may be absorbed by your sample, but red might not? The absorbance spectrum of the molybdenum-blue complex is very specific and you could use this to your advantage using spectral photometry.

Photometry + correcting for absorbance due to cell growth

Another approach is to quantify over time how your bacteria affect absorbance as a function of time and then subtract this from your experimental values. For example, in a wellplate, place the following adjacently: use a (i) well without cells, and only arsenite in solution, (ii) cells without arsenite, and (iii) cells with arsenite. This should allow you to quantify quite accurately how much the arsenite, the cells, and how much both affect absorbance.

Of course as you say there may be an interaction between the cells and the arsenite slowing or speeding up cell growth, so that would be a pickle to disentangle. You could try, perhaps in the wellplates (this would be best!), to find out what effect arsenite has on growth and correct your final data this way.

Periodically sampling the solution; removing bacteria from the measurement

Another idea would be to culture your cells in arsenite, and take samples to obtain time-points. For each sample, you filter using small syringe filters (these are cheap and are typically used for sterilizing solutions, i.e. getting rid of bacteria but keeping the solution, given their pore size of ~.1 microns) and then perform photometry for the molybdenum-blue complex. This is typical practice in microbiology.

PCR? An idea.

Last but not least, it's good to remember this is an indirect readout of transgenic gene expression. Similarly, you could use quantitative PCR to quantify the expression of any transgenes - here, you use housekeeping genes as controls for cell number, and the mRNA expression of your transgene as the readout. You would be reading out the expression of mRNA, but I personally don't think it would be more 'indirect' than measuring arsenate concentrations in solution, especially when there are confounding factors and awkwardness in getting a clean measurement. The PCR approach would be quite accurate and likely more reliable.

These are just some initial thoughts - perhaps you could add a comment or we could chat about this if you think it could spark up some concrete ideas!


Following Abeta fibrillization/aggregation in real-time using a FLUOstar Omega microplate reader

Aggregation of the amyloid-ß (Aß) peptide is a fundamental hall-mark for Alzheimer’s disease. The formation of extracellular senile plaques will lead to synaptic and neuronal damages in clinical demented patients. The aggregation process of Aß peptide is seen as seed driven. These seeds consist of small stable aggegrates of Aß. It is thought that these aggregates are already present in early stages of Alzheimer’s even before a patient experiences any symptoms. If this is true, determination of these early aggregates (aggregation seeds) would be an excellent diagnostic tool.


Here we present a cell-free assay (FRANK-Assay = Fibrilization of recombinant Aß nucleation kinetics) that allows determination of the amount of aggregation seeds from brain tissue homogenates. The assay is run over 2-3 days using the FLUOstar ® Omega microplate reader from BMG LABTECH.


Introduction

Genetic modification of crop plants often has the goal to engineer lines that express novel traits that cannot be introduced into the crop by conventional breeding. Such bioengineering efforts build on the expectation that target gene(s) conferring the desired trait, in association with suitable regulatory elements that are also part of the transgene construct, express the desired trait in a stable and reliable manner. This expectation remains to be evaluated, for example when the transgene is introduced into different genetic background (i.e. varieties) or when genetically modified (GM) plants are exposed to diverse environmental conditions.

One of the two most widely marketed GM traits worldwide is insect resistance, which is conferred by insecticidal toxins from Bacillus thuringiensis (Bt). This trait has been engineered into a number of crop plants, including maize and cotton. Maize containing insect resistance trait is currently grown on more than 47 million hectares worldwide [1], which represents about 27% of the global area planted with maize [2]. Bt maize cultivars derived from the MON810 event specifically contain a transgene cassette consisting of the cauliflower mosaic virus 35S promoter, thet e hsp70 intron and the cry1Ab gene endowing the resulting MON810 Bt maize plants with a resistance to lepidopteran pest species, particularly, the European corn borer, Ostrinia nubilalis.

It is generally expected that in commercial GM plants, transgenes are constitutively expressed at high levels and in all plant tissues and phenological phases [3]. Tight control over transgene expression and Bt protein content is important in light of concerns over the evolution of Bt toxin resistance in target insects [4]. In recent years, several studies have reported that the Bt protein concentration may vary within Bt plants (i.e. across tissues) and over growing seasons [5–7], while other authors reported that abiotic factors, such as nitrogen fertilization [8], soil quality and pesticide use [9] can affect Bt protein content. However, to the best of our knowledge, no study has been published to date that jointly investigated the relationship between Bt transgene expression and Bt protein content in transgenic Bt maize. In Bt cotton, this relationship has been investigated in a limited number of studies [10–12]. Olsen et al. [12] and Adamczyk et al. [10] found correlations between mRNA transcript levels of insect resistance transgene cry1Ac and Bt protein content, whereas Li et al. [11] observed no such relationship under salt stress. It therefore currently remains open what the relationship between Bt transgene expression and Bt protein content in GM crops is.

Establishing whether such a relationship exists in Bt maize and how it is affected by environmental conditions is an important question. In most countries where cultivation of Bt maize has been approved, this was done on the condition of installing an insect resistance management (IRM) program. One of the pillars of IRM is that plants contain high and stable levels of the Bt protein that are lethal not only to susceptible target insects but also to heterozygotes that carry one resistance allele (RS genotypes) [13]. The aim of IRM is to delay the evolution of resistance to Bt toxins in target pests which has been identified as a prime threat to the sustainability of Bt crops [4].

The aims of this study were to explore the relationship between Bt transgene expression and Bt protein content in two Bt maize varieties, and to experimentally test whether abiotic environmental stress conditions influence the relationship between transgene expression and protein content.


1 INTRODUCTION

The middle silk gland (MSG) and posterior silk gland (PSG) of silkworms, Bombyx mori, produce large quantities of silk thread proteins. 1 Recombinant versions of proteins produced in these MSGs 1-4 and PSGs 5-8 are used in protein engineering. The PSG expresses the biopolymer protein fibroin, which is highly biodegradable, biocompatible, and shows low immunostimulatory activity, making it suitable to produce surgical sutures 5 and grafts for revascularization, 9 bone tissue regeneration, 10 and cutaneous wound healing. 11, 12

Posterior silk glands engineered to express cell growth factors could be processed into biomedical materials, such as powders with high concentrations of fibroin, which could control the proliferation of cells in vivo and in vitro. However, one potential disadvantage of this system is that cell growth factors, including the human fibroblast growth factor-7 (FGF-7), are typically unstable under harsh environmental conditions, causing them to have a short half-life. 13, 14

The B.mori cypovirus 1 (CPV), which is a member of the family Reoviridae, produces proteinaceous occlusion bodies known as polyhedra that include microcrystals of the protein polyhedrin, which has attracted attention for its potential ability to encapsulate and protect proteins. 15-18 This virus infects cells in the silkworm digestive tract, producing large numbers of polyhedra. Progeny viruses are occluded in polyhedra that protect against infectivity over the long term and in the outdoor environment. 15 We previously showed that polyhedra could encapsulate diverse foreign proteins, such as fluorescence proteins, 16-19 cytokines, 20-23 and fusion proteins comprising enzymes 24 with an N-terminal polyhedron-encapsulation signal alpha-helix sequence H1 and C-terminal VP3-tag that are expressed during polyhedron crystallization in cultured insect cells. In addition, the cytokine activities in polyhedra are stable in the long term. 16 The remarkable stability of polyhedra-encapsulated proteins suggests that these systems could be robust as sustained-release carriers of cytokines and other proteins for tissue engineering. 16, 20-24

We also recently reported that the introduction of polyhedra into an in vitro neurodifferentiation cell culture model induced the release of active neurotrophin from polyhedra upon gradual polyhedra proteolysis mediated by small amounts of cell-derived matrix metalloproteinases. 23 These findings highlight the potential biomedical applications of polyhedra that encapsulate cytokines, such as FGF-7, inside silkworm silk glands. These silk glands can be processed to yield silk materials incorporating cytokine-polyhedra to control cell proliferation.

Our earlier studies showed that protein expression systems involving bioengineered silk glands could effectively produce active cytokines, such as fibroblast growth factor-2 (FGF-2 ref. 7). Here, we focused on the generation of transgenic silkworms using PSGs and MSGs that produce polyhedron-encapsulated FGF-7. FGF-7 is used for establishment of a three-dimensional (3D) culture system that serves as an in vitro model epidermis. 25 We examined whether the biological activity of FGF-7 can be released from the polyhedra to induce keratinocyte proliferation and epidermal differentiation of cells in supplement-free culture. We also investigated the effectiveness of FGF-7 activity released from polyhedra that are incorporated in PSGs in 3D keratinocyte cultures to inform the construction of a human epidermis model.


Introduction

Fluorescent protein (FP) technology has revolutionized many fields in the life sciences, including molecular biology, cell biology, biomedicine and biotechnology. Indeed, the 2008 Nobel Prize in Chemistry was awarded for discovery and development of the Aequorea victoria green fluorescent protein (avGFP, Chalfie et al., 1994 Heim et al., 1995 Shimomura et al., 1962 ). Today, FP technology is being used in an ever-expanding list of applications, including investigation of promoter function, protein localization, intracellular protein dynamics, protein–protein interactions, cell-cycle progression, organelle and organ labeling, transgenic organism tracking, and as biosensors to monitor small molecules and second messengers, just to name a few (Stewart, 2006 Shaner et al., 2007 Berg and Beachy, 2008 Day and Davidson, 2009 Frommer et al., 2009 Rizzo et al., 2009 ). Thus, FPs have become essential tools for biological research. However, such tools have not been widely developed for microalgal research.

Microalgae are single-celled photosynthetic organisms that have recently gained attention for their potential in biotechnology, including the production of biofuels, nutraceuticals such as omega-3 fatty acids, therapeutic proteins, and fish and animal feed (Pulz and Gross, 2004 Spolaore et al., 2006 Rosenberg et al., 2008 Radakovits et al., 2010 Specht et al., 2010 Yu et al., 2011 Georgianna and Mayfield, 2012 ). Chlamydomonas reinhardtii is a freshwater, green microalga that has become a popular model organism for photosynthesis and biotechnological research (Harris et al., 2009 ). C. reinhardtii has also shown promise as a production platform for human and animal therapeutic proteins and industrial enzymes (e.g.(Dreesen et al., 2010 Eichler-Stahlberg et al., 2009 Gregory et al., 2012 He et al., 2007 Jones et al., 2013 Manuell et al., 2007 Mayfield et al., 2003 Rasala et al., 2012 , 2010 Sun et al., 2003 Surzycki et al., 2009 Tran et al., 2012 , 2009 Wang et al., 2008 Yang et al., 2006 Zhang et al., 2006 ).

As both a model alga and a potential industrial strain, Chlamydomonas research would greatly benefit from development of FP tools and technology. Currently, only one FP, GFP, has been widely used in Chlamydomonas research. A nuclear codon-optimized gene has been developed for expression from the nuclear genome (CrGFP, Fuhrmann et al., 1999 ), and a chloroplast codon-optimized gene has been developed for expression from the chloroplast genome (Franklin et al., 2002 ). Although expression of GFP itself or as a chimeric protein in the chloroplast offers several advantages over nuclear expression, including higher levels of protein accumulation, which facilitates fluorescence detection and therefore utility, the disadvantage of chloroplast-expressed GFP is that it remains confined to the chloroplast. Therefore, many FP applications are not available with this method, including tagging of proteins not localized to the chloroplast, organelle labeling, and nuclear promoter studies, among many others. However, GFP expression from the nuclear genome has been severely hindered by low levels of protein expression, which has been attributed to the robust gene silencing mechanism(s) in Chlamydomonas (Fuhrmann et al., 1999 Neupert et al., 2009 ). The problem of poor GFP expression is compounded by the fact that Chlamydomonas produces a variety of highly fluorescent pigments, such as chlorophylls and flavonoids. Thus, poor transgene expression from the nuclear genome together with high auto-fluorescence has impeded the use of FPs in microalgae.

While engineering the nuclear genome for robust expression of monomeric GFP has proven challenging, some success has been achieved with GFP fusion proteins. GFP has been fused to the antibiotic resistance gene sh-ble (Stevens et al., 1996 ) and shown to localize to the nucleus (Fuhrmann et al., 1999 Rasala et al., 2012 ). sh-ble confers antibiotic resistance through drug sequestration rather than enzymatic inactivation (Dumas et al., 1994 ), making it a good fusion partner because high levels of protein expression are required to allow cell survival during antibiotic selection (Fuhrmann et al., 1999 Rasala et al., 2012 ). GFP has also been fused to endogenous genes, but with mixed results. For example, GFP has been fused to flagella proteins and successfully imaged by fluorescence microscopy in live cells (e.g. Schoppmeier et al., 2005 Huang et al., 2007 Diener, 2009 Engel et al., 2009 ). However, this often requires specialized live-cell imaging techniques in order to detect the GFP signal over the overwhelming auto-fluorescence generated by the cell body. Techniques include using a mosaic digital illumination system (Photonic Instruments, St. Charles, IL, USA) to control the area of the specimen that is illuminated, or confocal or total internal reflection fluorescence microscopy, which requires adherence of the flagella to a cover slip, with the auto-fluorescent cell bodies positioned outside the field of illumination (Diener, 2009 ). GFP has also been fused to proteins of the cell body. However, to overcome auto-fluorescence, visualization of the plasma membrane protein Rh1 fused to GFP, for example, required expression from a white mutant strain of Chlamydomonas, which is blocked at the first step of carotenoid biosynthesis (Yoshihara et al., 2008 ).

To our knowledge, the only other FP that has been expressed in Chlamydomonas to date is YFP (Neupert et al., 2009 ). In that study, the authors used UV mutagenesis to create strains of Chlamydomonas that express detectable levels of heterologous proteins from the nuclear genome, including monomeric YFP and GFP.

The aim of the present study was to express a full complement of FPs that span the visual spectrum in the green microalga Chlamydomonas, and to evaluate and compare their performance as reporter genes. To overcome the challenge of poor transgene expression from the nuclear genome, we fused the FPs to the sh-ble selection marker (Ble). To enable accumulation of unfused, untargeted and monomeric FPs, we inserted the self-cleaving 2A peptide from the foot and mouth disease virus (FMDV Ryan et al., 1991 ) between Ble and the FPs. We have previously shown that the FMDV 2A peptide efficiently self-cleaves in Chlamydomonas, and that the ble–2A expression strategy led to high levels of CrGFP accumulation, approximately 0.25% of total soluble protein, and detectable fluorescence in live-cell microscopy (Rasala et al., 2012 ). In this study, we show successful expression of five additional FPs that span the visual spectrum (blue mTagBFP, cyan mCerulean, yellow Venus, orange tdTomato and red mCherry), and compare them to green CrGFP. All FPs were detectable by immunoblotting, fluorescence microplate reader analysis on whole cells, fluorescence microscopy, and flow cytometry. Interestingly, CrGFP was shown to be the least fluorescent due to its low signal-to-noise ratio, while the FPs with longer emission wavelengths (Venus, tdTomato and mCherry) had the highest signal-to-noise ratios. Finally, we show that the ble–2A expression vector may be used to tag an endogenous gene, α-tubulin, with a fluorescent protein tag that is readily detectable using standard live-cell imaging techniques.


Results and Discussion

To establish a method for multiple measurements of gene expression in the same embryos, we exploited properties of GFP that are amenable for quantification namely, it diffuses freely in the cytoplasm, it is readily detected, and the fluorescence signal is proportional to the amount of protein. We noted that following injection of rGFP into the cytoplasm of 1-cell embryos 1) the rGFP rapidly dispersed throughout the cytoplasm, 2) as little as 3 pg of protein could be detected above background autofluorescence, and 3) that a highly reproducible and linear signal was observed over a range of 3–60 pg of protein, as determined by computer analysis of photo images of the fluorescing embryos ( Fig. 1B). It should be noted that because rGFP is relatively refractory to proteolysis [ 18], the fluorescence intensity was stable for at least 3 days when these rGFP-injected embryos were stored at 4°C. This permitted the use of a single set of injected embryos to serve as the GFP fluorescence standard for multiple experiments.

The vast majority of the pronuclear-injected embryos showed a nonmosaic and uniform green fluorescent signal ( Fig. 2). In some embryos, however, the signal was uneven among blastomeres and, considering that the bulk of transcription driven from the plasmid-borne reporter gene is from unintegrated copies, this could have been due to the unequal distribution of microinjected DNA among dividing blastomeres. If so, this occurred preferentially during the mid-preimplantation stages rather than during the first cleavage, because the unevenly fluorescing embryonic area (both stronger and weaker) was usually the minor area of the embryo.

Nonmosaic d1EGFP expression in the vast majority of the pronuclear-injected embryos (A, UV view B, bright light view). Some embryos show uneven green fluorescent signal as an indicator of apparent unequal distribution of microinjected DNA. Arrowheads indicate unevenly fluorescing embryonic areas (both stronger and weaker)

Nonmosaic d1EGFP expression in the vast majority of the pronuclear-injected embryos (A, UV view B, bright light view). Some embryos show uneven green fluorescent signal as an indicator of apparent unequal distribution of microinjected DNA. Arrowheads indicate unevenly fluorescing embryonic areas (both stronger and weaker)

To verify that expression of d1EGFP did not impair preimplantation development, we compared the developmental capacity of embryos expressing d1EGFP with the developmental capacity of embryos injected with buffer alone (sham). Analysis of the results of 4 experiments indicated that there was no significant difference in the developmental rate to the blastocyst stage between EGFP gene-injected embryos (74% 142 out of 192 embryos that survived injection) and sham-injected embryos (79% 38 out of 48 injected that survived injection). In previous studies, reduced developmental competence was observed among both the GFP-fluorescing and nonfluorescing embryos [ 22, 28, 29], suggesting that there is no direct link between reduced embryo viability and GFP/EGFP expression.

In order to monitor gene expression in the same embryo multiple times, it was necessary to establish conditions for UV irradiation that could detect the fluorescence but not inhibit development in vitro. Accordingly, we tested the influence of multiple UV exposures on embryo viability. Embryos were first exposed to UV irradiation at the two-cell stage followed by UV irradiation each day up to the blastocyst stage. Under our conditions of illumination, there was no apparent difference in the incidence of development to the blastocyst stage of these illuminated embryos to that of control embryos that were not exposed to UV light ( Table 1). More important, multiple exposures to UV light did not affect the developmental capacity of embryos. Longer (120 sec), multiple exposures to UV light decreased the incidence of development to the blastocyst stage, and no development to the blastocyst stage was observed after multiple (5–10) and prolonged (120 sec) exposures to UV light at the 2-cell stage.

In vitro development of mouse embryos a subjected to multiple UV irradiation exposure.

UV exposure . Blastocyst/observed embryo .
30 sec . 120 sec .
Single 43/54 (80%) 34/45 (76%)
Multiple/time course b 142/192 (74%) c 21/50 (42%)
Multiple/day d 9/50 (18%) 0/38 (0%)
Not exposed 29/36 (80%)
UV exposure . Blastocyst/observed embryo .
30 sec . 120 sec .
Single 43/54 (80%) 34/45 (76%)
Multiple/time course b 142/192 (74%) c 21/50 (42%)
Multiple/day d 9/50 (18%) 0/38 (0%)
Not exposed 29/36 (80%)

Injected at the 1-cell stage with pCMV-d1EGFP DNA.

One-two times a day, each day within the time course.

Data combined from 4 separated experiments.

Five-ten times a day, 1 day.

In vitro development of mouse embryos a subjected to multiple UV irradiation exposure.

UV exposure . Blastocyst/observed embryo .
30 sec . 120 sec .
Single 43/54 (80%) 34/45 (76%)
Multiple/time course b 142/192 (74%) c 21/50 (42%)
Multiple/day d 9/50 (18%) 0/38 (0%)
Not exposed 29/36 (80%)
UV exposure . Blastocyst/observed embryo .
30 sec . 120 sec .
Single 43/54 (80%) 34/45 (76%)
Multiple/time course b 142/192 (74%) c 21/50 (42%)
Multiple/day d 9/50 (18%) 0/38 (0%)
Not exposed 29/36 (80%)

Injected at the 1-cell stage with pCMV-d1EGFP DNA.

One-two times a day, each day within the time course.

Data combined from 4 separated experiments.

Five-ten times a day, 1 day.

We next quantified dlEGFP expression in single embryos in which multiple measurements were made on the same embryo ( Fig. 3). In these experiments the signal intensity present in the embryos was converted to the amount of protein, and the average dlEGFP content was calculated for groups of 20–30 embryos (a total of ∼100 embryos in 4 experiments) for each data point during the time-course experiment. Embryos arrested in development during culture were eliminated from analysis.

Quantitative analysis of the temporally restricted changes in transcription with the use of short half-life d1EGFP. Embryos were injected with 2 pl of DNA at both the 1-cell (squares) and 2-cell (circles) stages followed by culturing with (█, •) or without (□, ○) the presence of 1.5 mM butyrate. Multiple noninvasive measurements of d1EGFP expression were performed on the same embryos at various times after injection of DNA (10 ng/μl) to determine the dynamics in the steady state level of d1EGFP mRNA. Kinetics of d1EGFP expression were determined from the fluorescence values that were converted to the amount of d1EGFP. d1EGFP values observed for microinjected embryos were subtracted before calculating the mean ± SEM. Data from 4 independent experiments (total of ∼100 embryos) were combined, and the SEM was calculated

Quantitative analysis of the temporally restricted changes in transcription with the use of short half-life d1EGFP. Embryos were injected with 2 pl of DNA at both the 1-cell (squares) and 2-cell (circles) stages followed by culturing with (█, •) or without (□, ○) the presence of 1.5 mM butyrate. Multiple noninvasive measurements of d1EGFP expression were performed on the same embryos at various times after injection of DNA (10 ng/μl) to determine the dynamics in the steady state level of d1EGFP mRNA. Kinetics of d1EGFP expression were determined from the fluorescence values that were converted to the amount of d1EGFP. d1EGFP values observed for microinjected embryos were subtracted before calculating the mean ± SEM. Data from 4 independent experiments (total of ∼100 embryos) were combined, and the SEM was calculated

A green fluorescent signal was first detected in embryos 40 h after microinjection of the male pronucleus (i.e., at the 4-cell stage). Others have reported expression of a plasmid-borne luciferase reporter gene within 20–24 h following microinjection [ 1]. The delay that we observed may reflect a combination of a reduced sensitivity of detection of EGFP relative to luciferase and the small amount of plasmid DNA microinjected (0.02 pg), compared with the larger amounts (0.2–0.4 pg) injected in previous studies [ 1, 2]. Expression remained essentially constant up to 70 h following microinjection and then progressively declined until little if any expression was observed by 100 h postinjection (i.e., the blastocyst stage). This decrease likely reflected the degradation of the injected plasmid and not a decline in the transcriptional activity, because PCR analysis revealed a decrease in the amount of plasmid DNA ( Fig. 4). We observed that within the first hour after microinjection a 30%–50% decrease in the amount of plasmid DNA had occurred as suggested previously [ 30], this decrease is likely due to the leakage of the injected DNA into the cytoplasm, where it is rapidly degraded. By the expanded blastocyst stage (100 h postinjection) only ∼5% of the plasmid DNA remains.

Quantitative PCR analysis of the remaining plasmid DNA content at various times after pronuclear microinjection. A) One-cell embryos were injected with ∼4000 copies of the plasmid (which corresponds to 0.02 ng of the 4.9-kilobase plasmid) and PCR analysis was conducted on 10 embryos at different developmental stages. Shown is an ethidium bromide-stained 2% agarose gel. B) End-point titration standard. Lanes 1 to 8 show amplification fragments derived from 40 000, 20 000, 10 000, 5000, 3000, 2000, 1000, and 500 plasmid copies per embryo, respectively. Note that 40 000 copies of the plasmid is equivalent to the amount of DNA injected into 10 embryos

Quantitative PCR analysis of the remaining plasmid DNA content at various times after pronuclear microinjection. A) One-cell embryos were injected with ∼4000 copies of the plasmid (which corresponds to 0.02 ng of the 4.9-kilobase plasmid) and PCR analysis was conducted on 10 embryos at different developmental stages. Shown is an ethidium bromide-stained 2% agarose gel. B) End-point titration standard. Lanes 1 to 8 show amplification fragments derived from 40 000, 20 000, 10 000, 5000, 3000, 2000, 1000, and 500 plasmid copies per embryo, respectively. Note that 40 000 copies of the plasmid is equivalent to the amount of DNA injected into 10 embryos

Injection of the plasmid into the nucleus of a two-cell blastomere resulted in the rapid appearance of detectable fluoresence that was initially higher than that observed following injection of the male pronucleus ( Fig. 3). A rapid increase is also observed following injection of a 2-cell blastomere nucleus with the plasmid-borne luciferase reporter gene, and this level of expression is likewise greater than that observed following injection of 1-cell embryos [ 2]. Expression then increased and remained essentially constant for the next 40 h, after which it revealed a progressive decrease up to the blastocyst stage. Again, the decrease likely reflected the degradation of the injected plasmid, as determined by PCR analysis ( Fig. 4), and not a decline in transcriptional activity.

A transcriptionally repressive state develops during the maternal-to-zygotic transition ([ 31] and references therein). This repression has been detected by the requirement for an enhancer for high levels of expression of a plasmid-borne luciferase reporter gene following injection into a 2-cell nucleus [ 3]. Moreover, inducing histone hyperacetylation by butyrate treatment relieves this requirement. It is interesting that expression of the plasmid following microinjection into the male pronucleus shows neither a requirement for an enhancer nor increased expression following butyrate treatment [ 32]. Accordingly, we examined the effect of inducing histone hyperacetylation with butyrate on expression of our reporter gene following injection of either the male pronucleus or two-cell blastomere nucleus ( Fig. 3). Little stimulation of dlEGFP expression was observed in butyrate-treated 1-cell embryos, whereas a robust stimulation was found following injection of a 2-cell nucleus, when compared with embryos not treated with butyrate. Thus, monitoring dlEGFP fluorescence as a measure of reporter gene expression faithfully mimics the expression of a luciferase reporter gene and the effect of inducing histone hyperacetylation on reporter gene expression.

In summary, the method described in this report offers the investigator the opportunity to assay quantitatively in real-time the transcriptional activity of individual embryos under conditions that permit multiple measurements. The method should prove quite valuable in assessing the transcriptional activity in the preimplantation embryo (e.g., ascertaining the effect of different culture conditions on transcription, investigating the molecular mechanisms that underlie the maternal-to-zygotic transition, and analyzing transcription in species in which only a limited number of embryos are available [in nonhuman primates]).


RESULTS

Characterization of Transgenic Plants Expressing Polyprotein Constructs with IbAMP-Based Linker Peptides

A, Diagram of the expression cassettes in plant transformation vectors pFAJ3105, pFAJ3109, pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375. p35S, Enhanced CaMV35S promoter ( Kay et al., 1987) TMV, 5′ leader sequence of tobacco mosaic virus SP, coding region of the leader peptide derived from the DmAMP1 precursor DmAMP1, coding region of the mature DmAMP1 LP, coding region of the linker peptide RsAFP2, coding region of the mature RsAFP2 tNos, 3′-untranslated terminator region of the Agrobacterium tumefaciens nopaline synthase gene ( Bevan et al., 1983) [uOcs]pMas, chimeric promoter consisting of the enhancer of theA. tumefaciens octopine synthase gene and the promoter of the A. tumefaciens mannopine synthase gene (M.F.C. De Bolle, unpublished data) tMas, 3′-untranslated terminator region of theA. tumefaciens mannopine synthase gene ( Barker et al., 1983). B, Amino acid sequence of the (poly)proteins encoded by constructs pFAJ3105, pFAJ3109, pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375. The leader peptide of DmAMP1 is in normal font. The mature DmAMP1 domain is underlined. The linker peptide domain is in bold. The mature RsAFP2 domain is italicized.

A, Diagram of the expression cassettes in plant transformation vectors pFAJ3105, pFAJ3109, pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375. p35S, Enhanced CaMV35S promoter ( Kay et al., 1987) TMV, 5′ leader sequence of tobacco mosaic virus SP, coding region of the leader peptide derived from the DmAMP1 precursor DmAMP1, coding region of the mature DmAMP1 LP, coding region of the linker peptide RsAFP2, coding region of the mature RsAFP2 tNos, 3′-untranslated terminator region of the Agrobacterium tumefaciens nopaline synthase gene ( Bevan et al., 1983) [uOcs]pMas, chimeric promoter consisting of the enhancer of theA. tumefaciens octopine synthase gene and the promoter of the A. tumefaciens mannopine synthase gene (M.F.C. De Bolle, unpublished data) tMas, 3′-untranslated terminator region of theA. tumefaciens mannopine synthase gene ( Barker et al., 1983). B, Amino acid sequence of the (poly)proteins encoded by constructs pFAJ3105, pFAJ3109, pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375. The leader peptide of DmAMP1 is in normal font. The mature DmAMP1 domain is underlined. The linker peptide domain is in bold. The mature RsAFP2 domain is italicized.

Northern-blot analysis of DmAMP1 expression in leaves of transgenic T1 generation lines transformed with constructs pFAJ3109 and pFAJ3105. All analyzed samples represent 15 μg of total RNA. NT, Non-transformed Arabidopsis plants 3109, Arabidopsis plants transformed with the single-protein construct pFAJ3109 (transgenic line 1 and 2) 3105, Arabidopsis plants transformed with the double-protein construct pFAJ3105 (transgenic line 1 and 2). Similar northern-blot analyses were obtained for another eight lines of Arabidopsis plants transformed with construct pFAJ3105 and six lines of Arabidopsis plants transformed with construct pFAJ3109.

Northern-blot analysis of DmAMP1 expression in leaves of transgenic T1 generation lines transformed with constructs pFAJ3109 and pFAJ3105. All analyzed samples represent 15 μg of total RNA. NT, Non-transformed Arabidopsis plants 3109, Arabidopsis plants transformed with the single-protein construct pFAJ3109 (transgenic line 1 and 2) 3105, Arabidopsis plants transformed with the double-protein construct pFAJ3105 (transgenic line 1 and 2). Similar northern-blot analyses were obtained for another eight lines of Arabidopsis plants transformed with construct pFAJ3105 and six lines of Arabidopsis plants transformed with construct pFAJ3109.

The amount of DmAMP1 and RsAFP2 in leaves from a series of T1 Arabidopsis plants transformed with pFAJ3105 was determined using ELISA assays (Table I). Most of the lines transformed with the polyprotein construct expressed both DmAMP1-cross-reactive proteins (CRPs) and RsAFP2-CRPs. In general, there was a good correlation between DmAMP1-CRP and RsAFP2-CRP levels. However, RsAFP2-CRP levels were generally 2- to 5-fold lower than the DmAMP1-CRP levels. The ELISA assays for measuring the RsAFP2-CRP levels in the extracts were, however, less reliable than those for the DmAMP1-CRPs. In the RsAFP2 ELISA assays, dilutions of extracts of transgenic plants yielded dose-response curves that deviated from those obtained for dilutions of standard solutions containing native RsAFP2, indicating that the majority of the RsAFP2-CRPs in the extracts was not immunologically identical to the native RsAFP2 itself.

Expression levels of DmAMP1 and RsAFP2 in leaves of transgenic Arabidopsis transformed with the constructs pFAJ3105 and pFAJ3109

Construct . Line . Expression Level of DmAMP1 . Expression Level of RsAFP2 .
%
pFAJ3105 1 0.77 0.29
2 1.13 0.22
3 0.48 0.20
4 0.005 <0.001
5 0.36 0.05
6 0.99 0.25
7 0.60 0.09
8 0.13 <0.001
9 0.25 0.08
10 1.35 0.35
11 0.24 0.07
12 1.18 0.24
13 0.68 0.17
14 0.49 0.07
Average 0.62 0.15
pFAJ3109 1 0.19 nd 1-a
2 0.05 nd
3 0.02 nd
4 0.20 nd
5 0.10 nd
6 0.06 nd
7 0.07 nd
8 0.003 nd
Average 0.09 nd
Construct . Line . Expression Level of DmAMP1 . Expression Level of RsAFP2 .
%
pFAJ3105 1 0.77 0.29
2 1.13 0.22
3 0.48 0.20
4 0.005 <0.001
5 0.36 0.05
6 0.99 0.25
7 0.60 0.09
8 0.13 <0.001
9 0.25 0.08
10 1.35 0.35
11 0.24 0.07
12 1.18 0.24
13 0.68 0.17
14 0.49 0.07
Average 0.62 0.15
pFAJ3109 1 0.19 nd 1-a
2 0.05 nd
3 0.02 nd
4 0.20 nd
5 0.10 nd
6 0.06 nd
7 0.07 nd
8 0.003 nd
Average 0.09 nd

Expression levels of DmAMP1 and RsAFP2 are expressed as the ratio of the amount of DmAMP1-CRP or RsAFP2-CRP to the amount of total soluble protein in crude extracts from leaves of transgenic Arabidopsis plants. The Arabidopsis plants are transformed with either the double protein construct (pFAJ3105) coding for DmAMP1 and RsAFP2 or the single protein construct (pFAJ3109) coding only for DmAMP1.

Expression levels of DmAMP1 and RsAFP2 in leaves of transgenic Arabidopsis transformed with the constructs pFAJ3105 and pFAJ3109

Construct . Line . Expression Level of DmAMP1 . Expression Level of RsAFP2 .
%
pFAJ3105 1 0.77 0.29
2 1.13 0.22
3 0.48 0.20
4 0.005 <0.001
5 0.36 0.05
6 0.99 0.25
7 0.60 0.09
8 0.13 <0.001
9 0.25 0.08
10 1.35 0.35
11 0.24 0.07
12 1.18 0.24
13 0.68 0.17
14 0.49 0.07
Average 0.62 0.15
pFAJ3109 1 0.19 nd 1-a
2 0.05 nd
3 0.02 nd
4 0.20 nd
5 0.10 nd
6 0.06 nd
7 0.07 nd
8 0.003 nd
Average 0.09 nd
Construct . Line . Expression Level of DmAMP1 . Expression Level of RsAFP2 .
%
pFAJ3105 1 0.77 0.29
2 1.13 0.22
3 0.48 0.20
4 0.005 <0.001
5 0.36 0.05
6 0.99 0.25
7 0.60 0.09
8 0.13 <0.001
9 0.25 0.08
10 1.35 0.35
11 0.24 0.07
12 1.18 0.24
13 0.68 0.17
14 0.49 0.07
Average 0.62 0.15
pFAJ3109 1 0.19 nd 1-a
2 0.05 nd
3 0.02 nd
4 0.20 nd
5 0.10 nd
6 0.06 nd
7 0.07 nd
8 0.003 nd
Average 0.09 nd

Expression levels of DmAMP1 and RsAFP2 are expressed as the ratio of the amount of DmAMP1-CRP or RsAFP2-CRP to the amount of total soluble protein in crude extracts from leaves of transgenic Arabidopsis plants. The Arabidopsis plants are transformed with either the double protein construct (pFAJ3105) coding for DmAMP1 and RsAFP2 or the single protein construct (pFAJ3109) coding only for DmAMP1.

Box plots of the DmAMP1 expression levels of T1 generation transgenic Arabidopsis plants transformed with constructs pFAJ3105 and pFAJ3109, respectively. DmAMP1 expression level is expressed as the ratio of the amount of DmAMP1 to the amount of total soluble protein in crude extracts from leaves of transgenic T1 Arabidopsis plants. Gridlines depicted as vertical dotted lines represent the DmAMP1 expression level scale. Vertical lines in the box plots correspond to the 0th, 25th, 50th (or median), 75th, and 100th percentile, respectively.

Box plots of the DmAMP1 expression levels of T1 generation transgenic Arabidopsis plants transformed with constructs pFAJ3105 and pFAJ3109, respectively. DmAMP1 expression level is expressed as the ratio of the amount of DmAMP1 to the amount of total soluble protein in crude extracts from leaves of transgenic T1 Arabidopsis plants. Gridlines depicted as vertical dotted lines represent the DmAMP1 expression level scale. Vertical lines in the box plots correspond to the 0th, 25th, 50th (or median), 75th, and 100th percentile, respectively.

To test whether enhanced expression observed for the polyprotein construct pFAJ3105 is restricted to the particular configuration of the expression unit, four other constructs (pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375) were made (Fig. 1). The promoter used in these constructs was the chimeric promoter [uOcs]pMas, consisting of the A. tumefaciens octopine synthase enhancer linked to the A. tumefaciens mannopine synthase promoter instead of the CaMV35S promoter used in construct pFAJ3105. The terminator was the mannopine synthase terminator instead of the nopaline synthase terminator used in construct pFAJ3105. Constructs pFAJ3339 and pFAJ3340 are both polyprotein-encoding constructs, with the polyprotein encoded by pFAJ3339 being exactly the same as the polyprotein encoded by pFAJ3105 and the one encoded by pFAJ3340 having a reversed order of the DmAMP1 and RsAFP2 coding regions relative to that of pFAJ3105 and pFAJ3339. Constructs pFAJ3433 and pFAJ3375 are both single-protein constructs, with pFAJ3433 encoding mature DmAMP1 preceded by the DmAMP1 leader peptide at its amino terminus and pFAJ3375 encoding mature RsAFP2 with the DmAMP1 leader peptide.

Expression levels of DmAMP1 and RsAFP2 in leaves of transgenic Arabidopsis transformed with the constructs pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375

Construct . n 2-a . Average Expression Level . P 2-b . P 2-c .
DmAMP1 . RsAFP2 .
%
pFAJ3339 13 0.085 0.024 a a
pFAJ3340 19 0.006 0.01 b a
pFAJ3433 20 0.009 nd 2-d b nd
pFAJ3375 13 nd 0.003 nd b
Construct . n 2-a . Average Expression Level . P 2-b . P 2-c .
DmAMP1 . RsAFP2 .
%
pFAJ3339 13 0.085 0.024 a a
pFAJ3340 19 0.006 0.01 b a
pFAJ3433 20 0.009 nd 2-d b nd
pFAJ3375 13 nd 0.003 nd b

Expression levels are expressed as described in Table I.

n is the no. of independent transformants for which expression was measured.

Comparison of the average of the ln-transformed DmAMP1 expression levels at significance level 0.05. Identical letters indicate that the averages are not significantly different according to Tukey's studentized range test.

Comparison of the average of the ln-transformed RsAFP2 expression levels at significance level 0.05. Identical letters indicate that the averages are not significantly different according to Tukey's studentized range test.

Expression levels of DmAMP1 and RsAFP2 in leaves of transgenic Arabidopsis transformed with the constructs pFAJ3339, pFAJ3340, pFAJ3433, and pFAJ3375

Construct . n 2-a . Average Expression Level . P 2-b . P 2-c .
DmAMP1 . RsAFP2 .
%
pFAJ3339 13 0.085 0.024 a a
pFAJ3340 19 0.006 0.01 b a
pFAJ3433 20 0.009 nd 2-d b nd
pFAJ3375 13 nd 0.003 nd b
Construct . n 2-a . Average Expression Level . P 2-b . P 2-c .
DmAMP1 . RsAFP2 .
%
pFAJ3339 13 0.085 0.024 a a
pFAJ3340 19 0.006 0.01 b a
pFAJ3433 20 0.009 nd 2-d b nd
pFAJ3375 13 nd 0.003 nd b

Expression levels are expressed as described in Table I.

n is the no. of independent transformants for which expression was measured.

Comparison of the average of the ln-transformed DmAMP1 expression levels at significance level 0.05. Identical letters indicate that the averages are not significantly different according to Tukey's studentized range test.

Comparison of the average of the ln-transformed RsAFP2 expression levels at significance level 0.05. Identical letters indicate that the averages are not significantly different according to Tukey's studentized range test.

A, Box plots of the DmAMP1 expression levels of T1 generation Arabidopsis plants transformed with the polyprotein constructs pFAJ3339 and pFAJ3340 and the single-protein construct pFAJ3433, respectively. B, Box plots of the RsAFP2 expression levels of T1 generation Arabidopsis plants transformed with the polyprotein constructs pFAJ3339 and pFAJ3340 and the single-protein construct pFAJ33775, respectively. Legend for both box plots as in Figure 3.

A, Box plots of the DmAMP1 expression levels of T1 generation Arabidopsis plants transformed with the polyprotein constructs pFAJ3339 and pFAJ3340 and the single-protein construct pFAJ3433, respectively. B, Box plots of the RsAFP2 expression levels of T1 generation Arabidopsis plants transformed with the polyprotein constructs pFAJ3339 and pFAJ3340 and the single-protein construct pFAJ33775, respectively. Legend for both box plots as in Figure 3.

Analysis of the Subcellular Location of Co-Expressed Plant Defensins

To determine whether the co-expressed plant defensins are either secreted extracellularly or deposited intracellularly, extracellular fluid and intracellular extract fractions were prepared from leaves of Arabidopsis plants transformed with the polyprotein construct (pFAJ3105). The cytosolic enzyme Glc-6-phosphate dehydrogenase was used as a marker to detect contamination of the extracellular fluid fraction with intracellular components. Glc-6-phosphate dehydrogenase was partitioned in a ratio of 83:17 between the intracellular extract fractions and extracellular fluid fractions. In contrast, 94% of DmAMP1-CRP content and 92% of RsAFP2-CRP content in the transgenic plants tested were found in the extracellular fluid fractions. These results indicate that both plant defensins released from the polyprotein precursors are deposited primarily in the apoplast.

Purification of Proteins Processed from Polyprotein Construct pFAJ3105

Chromatographic purification of DmAMP1-CRPs and RsAFP2-CRPs from extracellular fluid fraction of Arabidopsis plants transformed with the double-protein construct pFAJ3105. The extracellular fluid fraction was loaded on a C8 RP-HPLC column equilibrated in 15% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid (TFA). The column was eluted at a flow rate of 1 mL min −1 for 15 min with 15% (v/v) acetonitrile in 0.1% (v/v) TFA followed by a linear gradient of acetonitrile in 0.1% (v/v) TFA from 15% (v/v) to 50% (v/v) acetonitrile in 35 min. The eluate was monitored by on-line measurement of theA 280 (A 280—) and the acetonitrile gradient (---) was monitored with an on-line conductivity sensor. Fractions were collected and assessed for the presence of DmAMP1-CRPs and RsAFP2-CRPs by the respective ELISA assays. Dotted bars represent the presence of DmAMP1-CRPs (p3105EF1 and p3105EF2), whereas the black bar represents the presence of RsAFP2-CRPs (p3105EF3). Triangles indicate the elution position of native DmAMP1 (white triangle) and of native RsAFP2 (black triangle).

Chromatographic purification of DmAMP1-CRPs and RsAFP2-CRPs from extracellular fluid fraction of Arabidopsis plants transformed with the double-protein construct pFAJ3105. The extracellular fluid fraction was loaded on a C8 RP-HPLC column equilibrated in 15% (v/v) acetonitrile in 0.1% (v/v) trifluoroacetic acid (TFA). The column was eluted at a flow rate of 1 mL min −1 for 15 min with 15% (v/v) acetonitrile in 0.1% (v/v) TFA followed by a linear gradient of acetonitrile in 0.1% (v/v) TFA from 15% (v/v) to 50% (v/v) acetonitrile in 35 min. The eluate was monitored by on-line measurement of theA 280 (A 280—) and the acetonitrile gradient (---) was monitored with an on-line conductivity sensor. Fractions were collected and assessed for the presence of DmAMP1-CRPs and RsAFP2-CRPs by the respective ELISA assays. Dotted bars represent the presence of DmAMP1-CRPs (p3105EF1 and p3105EF2), whereas the black bar represents the presence of RsAFP2-CRPs (p3105EF3). Triangles indicate the elution position of native DmAMP1 (white triangle) and of native RsAFP2 (black triangle).

To test whether the purification procedure based on the extracellular fluid preparation reflects the true composition in DmAMP1-CRPs and RsAFP2-CRPs of the transgenic Arabidopsis leaves, an alternative purification procedure was developed starting from a crude leaf extract. The extract was fractionated by ion-exchange chromatography (IEC) and subsequently by RP-HPLC. After separation, fractions were collected and assessed for the presence of DmAMP1-CRPs and RsAFP2-CRPs using ELISA assays. IEC was performed by passing the extract over a cation exchange column at pH 6. When the column was eluted with a linear gradient of 0 to 0.5 m NaCl in 50 m m MES at pH 6, DmAMP1-CRPs were detected in fractions eluting between 0.17 and 0.33 m NaCl, whereas RsAFP2-CRPs were detected in fractions eluting between 0.24 and 0.49 m NaCl (data not shown). Fractions containing either DmAMP1-CRPs or RsAFP2-CRPs were pooled into two fractions (0.17–0.24 m NaCl and 0.33–0.49 m NaCl, respectively), which were each subjected to RP-HPLC on a C8-silica column eluted with a linear gradient of acetonitrile. DmAMP1-CRPs eluted in two peaks, the latter of which eluted at a position very close to that of native DmAMP1. RsAFP2-CRPs were found in a single peak that was well separated from the DmAMP1-CRP peaks and eluted at a position very close to that of native RsAFP2 (data not shown).

Molecular Analyses of the Purified Protein Fractions

The different DmAMP1-CRPs and RsAFP2-CRPs purified from the extracellular fluid were subjected to amino-terminal amino acid sequence analysis as well as to mass spectrometry. The carboxy-terminal amino acid was determined based on the best approximation of the predicted theoretical mass to the experimentally determined mass (Table III). The DmAMP1-CRPs, p3105EF1 and p3105EF2 (nomenclature as in Fig. 5), had masses that were consistent with the presence of a single additional Ser residue at their carboxy-terminal end as compared with native DmAMP1. However, whereas the mass of p3105EF2 corresponded exactly (within experimental error) to that calculated for a DmAMP1 derivative with an additional Ser (hereafter called DmAMP1+S) at its carboxy-terminal end, the mass of p3105EF1 was in excess by 4 D relative to the calculated mass for DmAMP1+S. Hence, this protein might be a DmAMP1+S derivative with partially reduced disulfide bridges. This hypothesis was confirmed by reduction of the disulfide bonds in p3105EF1 and p3105EF2 with excess dithiothreitol, followed by separation of the reduced forms by RP-HPLC. The reduced form of p3105EF1 and p3105EF2 eluted at exactly the same position, indicating that p3105EF1 only differs from p3105EF2 in its disulfide bond content (data not shown).

Molecular analysis of the purified AMP fractions

Protein Fraction . Molecular Mass Determined by: . Amino Acid Sequence .
Matrix-assisted laser-desorption ionization time of flight MS . Electrospray ionization-MS . Theoretical prediction . Determined amino terminal . Predicted carboxy terminal .
D
p3105EF1 5,614 ± 5 5,608.3 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF2 5,602 ± 5 5,604.9 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF3 6,223 ± 6 nd 3-a 6,225.15 DVEPG QKICYFPC
Protein Fraction . Molecular Mass Determined by: . Amino Acid Sequence .
Matrix-assisted laser-desorption ionization time of flight MS . Electrospray ionization-MS . Theoretical prediction . Determined amino terminal . Predicted carboxy terminal .
D
p3105EF1 5,614 ± 5 5,608.3 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF2 5,602 ± 5 5,604.9 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF3 6,223 ± 6 nd 3-a 6,225.15 DVEPG QKICYFPC

Molecular mass (D) of DmAMP1-CRPs and RsAFP2-CRPs purified as described in Fig. 3 was determined by matrix-assisted laser-desorption-ionization time of flight or electrospray ionization-mass spectrometry (MS) and amino-terminal sequence of the same components was determined by automated Edman degradation. Also shown are the predicted carboxy-terminal sequences that give best correspondence between experimental molecular mass and theoretical molecular mass. Amino acids of the mature DmAMP1 domain are underlined, amino acids of the linker peptide are in bold, and amino acids of the mature RsAFP2 domain are italicized.

Molecular analysis of the purified AMP fractions

Protein Fraction . Molecular Mass Determined by: . Amino Acid Sequence .
Matrix-assisted laser-desorption ionization time of flight MS . Electrospray ionization-MS . Theoretical prediction . Determined amino terminal . Predicted carboxy terminal .
D
p3105EF1 5,614 ± 5 5,608.3 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF2 5,602 ± 5 5,604.9 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF3 6,223 ± 6 nd 3-a 6,225.15 DVEPG QKICYFPC
Protein Fraction . Molecular Mass Determined by: . Amino Acid Sequence .
Matrix-assisted laser-desorption ionization time of flight MS . Electrospray ionization-MS . Theoretical prediction . Determined amino terminal . Predicted carboxy terminal .
D
p3105EF1 5,614 ± 5 5,608.3 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF2 5,602 ± 5 5,604.9 ± 1 5,604.25 ELCEKAS CYFNC S
p3105EF3 6,223 ± 6 nd 3-a 6,225.15 DVEPG QKICYFPC

Molecular mass (D) of DmAMP1-CRPs and RsAFP2-CRPs purified as described in Fig. 3 was determined by matrix-assisted laser-desorption-ionization time of flight or electrospray ionization-mass spectrometry (MS) and amino-terminal sequence of the same components was determined by automated Edman degradation. Also shown are the predicted carboxy-terminal sequences that give best correspondence between experimental molecular mass and theoretical molecular mass. Amino acids of the mature DmAMP1 domain are underlined, amino acids of the linker peptide are in bold, and amino acids of the mature RsAFP2 domain are italicized.

The RsAFP2-CRP fraction p3105EF3 represents an RsAFP2 derivative with the additional pentapeptide sequence DVEPG at its amino terminus, corresponding to the five carboxy-terminal amino acids of the linker peptide. This protein is further referred to as DVEPG+RsAFP2.

The different DmAMP1-CRPs and RsAFP2-CRPs purified from total leaf extract from plants transformed with construct pFAJ3105 were analyzed in the same way. The analyses indicated that the same three molecular species were present in the total leaf extract, namely DmAMP1+S, a reduced form of DmAMP1+S and DVEPG+RsAFP2 (data not shown).

In addition, DmAMP1-CRPs and RsAFP2-CRPs were also purified from extracellular fluid of plants transformed with construct pFAJ3340, the construct in which the order of DmAMP1 and RsAFP2 in the polyprotein was reversed. In this case, the molecular entities identified were authentic RsAFP2 including a pyro-Glu residue at its amino-terminus, RsAFP2 with an amino-terminal pyro-Glu and an additional carboxy-terminal Ser (RsAFP2+S), and finally DmAMP1 with an additional amino-terminal pentapeptide (DVEPG+DmAMP1 data not shown). In the plants transformed with polyprotein construct pFAJ3340, no reduced or partially reduced forms of either DmAMP1-CRPs or RsAFP2-CRPs could be detected (data not shown).

In Vitro Antifungal Activity of the Purified Proteins

The antifungal activity of the purified fractions from the extracellular fluid of plants transformed with construct pFAJ3105, containing the major processed products DmAMP1+S and DVEPG+RsAFP2, respectively, were assayed using Fusarium culmorum as a test fungus (Table IV). The specific antimicrobial activity, expressed as protein concentration required for 50% growth inhibition of the test organism, of purified DmAMP1+S appeared to be identical to that of native DmAMP1. The specific antimicrobial activity of purified DVEPG+RsAFP2, however, was about 2-fold lower relative to that of native RsAFP2. The slight drop in antimicrobial activity of DVEPG+RsAFP2 is most likely because of the presence of the five additional amino-terminal amino acids. Nevertheless, our data prove that processing of the polyprotein precursors in transgenic plants can result in the release of bioactive proteins.

Antifungal activity of the purified AMP fractions

Protein Fraction . Antifungal Activity (IC50) .
p3105EF2 3.2 ± 0.7
DmAMP1 2.8 ± 0.5
p3105EF3 3.2 ± 0.8
RsAFP2 1.7 ± 0.4
Protein Fraction . Antifungal Activity (IC50) .
p3105EF2 3.2 ± 0.7
DmAMP1 2.8 ± 0.5
p3105EF3 3.2 ± 0.8
RsAFP2 1.7 ± 0.4

Antifungal activity was determined as the concentration of protein (micrograms per milliliter) required to cause 50% growth inhibition of the fungus F. culmorum relative to a control culture. Data are given as means of triplicates with se .

Antifungal activity of the purified AMP fractions

Protein Fraction . Antifungal Activity (IC50) .
p3105EF2 3.2 ± 0.7
DmAMP1 2.8 ± 0.5
p3105EF3 3.2 ± 0.8
RsAFP2 1.7 ± 0.4
Protein Fraction . Antifungal Activity (IC50) .
p3105EF2 3.2 ± 0.7
DmAMP1 2.8 ± 0.5
p3105EF3 3.2 ± 0.8
RsAFP2 1.7 ± 0.4

Antifungal activity was determined as the concentration of protein (micrograms per milliliter) required to cause 50% growth inhibition of the fungus F. culmorum relative to a control culture. Data are given as means of triplicates with se .


A potent Cas9-derived gene activator for plant and mammalian cells

Overexpression of complementary DNA represents the most commonly used gain-of-function approach for interrogating gene functions and for manipulating biological traits. However, this approach is challenging and inefficient for multigene expression due to increased labour for cloning, limited vector capacity, requirement of multiple promoters and terminators, and variable transgene expression levels. Synthetic transcriptional activators provide a promising alternative strategy for gene activation by tethering an autonomous transcription activation domain (TAD) to an intended gene promoter at the endogenous genomic locus through a programmable DNA-binding module. Among the known custom DNA-binding modules, the nuclease-dead Streptococcus pyogenes Cas9 (dCas9) protein, which recognizes a specific DNA target through base pairing between a synthetic guide RNA and DNA, outperforms zinc-finger proteins and transcription activator-like effectors, both of which target through protein–DNA interactions 1 . Recently, three potent dCas9-based transcriptional activation systems, namely VPR, SAM and SunTag, have been developed for animal cells 2,3,4,5,6 . However, an efficient dCas9-based transcriptional activation platform is still lacking for plant cells 7,8,9 . Here, we developed a new potent dCas9–TAD, named dCas9–TV, through plant cell-based screens. dCas9–TV confers far stronger transcriptional activation of single or multiple target genes than the routinely used dCas9–VP64 activator in both plant and mammalian cells.

Among synthetic gene activators, dCas9–TADs potentially offer unparalleled simplicity and multiplexability compared with zinc-finger protein–TADs and transcription activator-like effector (TALE)–TADs because synthetic guide RNAs (sgRNAs) can be easily modified to achieve new targeting specificities, and dCas9 guided by multiple sgRNAs can simultaneously bind to several different target loci 10 . However, a dCas9 fusion with VP64, a frequently used TAD 11 , only weakly activates target genes using a single sgRNA in plant and mammalian cells 7,8,9,12,13,14,15 . Using Arabidopsis protoplast-based promoter–luciferase (LUC) assays, we confirmed that dCas9–VP64 with a single sgRNA only weakly (maximally 2.4-fold) or ineffectively activated target genes (Supplementary Results and Supplementary Figs. 1 and 2). Interestingly, when the target sequence lacks a 5′ G, an extra G appended to the 5′ end of the sgRNA was found to enhance the promoter activation (Supplementary Fig. 1), presumably by promoting the transcription initiation of the sgRNA by the U6 promoter. Therefore, we routinely add a G to the 5′ end of sgRNAs when the target sequences start with a non-G nucleotide.


Overexpression of the Polygonum cuspidatum PcDREB2A Gene Encoding a DRE-Binding Transcription Factor Enhances the Drought Tolerance of Transgenic Arabidopsis thaliana

Plants have evolved complex signaling networks that enable them to adapt to adverse environmental conditions. The dehydration-responsive element-binding (DREB) transcription factors are important for plant responses to abiotic stresses. In this study, a new member of the AP2/ERF transcription factor gene family, PcDREB2A, was cloned and characterized from Polygonum cuspidatum, a traditional Chinese medicinal herb. PcDREB2A, which includes a typical AP2 domain, was clustered in the A-2 subgroup of the DREB subfamily. At the seedling stage, PcDREB2A expression was induced by cold, salt, and drought stresses. A yeast one-hybrid assay and an analysis of transiently transformed tobacco revealed that PcDREB2A can specifically bind to the DRE motif and transactivate reporter gene expression. Following 200 and 250 mM mannitol treatments, the PcDREB2A-overexpressing Arabidopsis thaliana lines had longer roots and a significantly higher fresh weight than the wild-type plants. Furthermore, under drought stress conditions, the PcDREB2A-overexpressing A. thaliana plants accumulated less malondialdehyde than the control plants. These results indicate that PcDREB2A encodes a novel DREB transcription factor in P. cuspidatum. Furthermore, the data generated in this study may be useful for researchers and breeders interested in genetically engineering plants to increase drought tolerance without inhibiting growth.

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Improved and high throughput quantitative measurements of weak GFP expression in transgenic plant materials

Green fluorescent proteins (GFPs) are widely used in tracing transgene expression and have been known as convenient and efficient markers for plant transformation. However, sometimes researchers are still puzzled by the weak fluorescence since it makes the observation of GFP signals and confirmation of transgenic plants difficult. In this investigation, we explored the possibility of enhancing the weak signals by changing the pH environment of detection and took microplate reader as a more effective instrument compared to traditional fluorescent microscope to detect the weak signals. It was found that the fluorescence intensity of enhanced GFP (EGFP) in transgenic plants can be increased 2–6 folds by altering the environmental pH, and the concentration of EGFP at a large scale (ranged from 20 ng/ml to 20 μg/ml) can be detected and quantified. It can exclude the influence of degradation fragment and hence facilitate later analysis these advantages were further verified by comparing with western blotting and confocal microscopy. It was reliable and effective for the qualitative and quantitative analysis of transgenic plants and was more suitable for the detection of very weak fluorescent signals.

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Watch the video: FlexStation 3 Multi-Mode Microplate Reader (December 2021).