Genetic Engineering of Dictyostelium discoideum Cells Based on Selection and Growth on Bacteria | Protocol

Jan, 25th 2019, 21:00 by

Dictyostelium discoideum is an intriguing model organism for the study of cell differentiation processes during development, cell signaling, and other important cellular biology questions. The technologies available to genetically manipulate Dictyostelium cells are well-developed. Transfections can be performed using different selectable markers and marker re-cycling, including homologous recombination and insertional mutagenesis. This is supported by a well-annotated genome. However, these approaches are optimized for axenic cell lines growing in liquid cultures and are difficult to apply to non-axenic wild-type cells, which feed only on bacteria. The mutations that are present in axenic strains disturb Ras signaling, causing excessive macropinocytosis required for feeding, and impair cell migration, which confounds the interpretation of signal transduction and chemotaxis experiments in those strains. Earlier attempts to genetically manipulate non-axenic cells have lacked efficiency and required complex experimental procedures. We have developed a simple transfection protocol that, for the first time, overcomes these limitations. Those series of large improvements to Dictyostelium molecular genetics allow wild-type cells to be manipulated as easily as standard laboratory strains. In addition to the advantages for studying uncorrupted signaling and motility processes, mutants that disrupt macropinocytosis-based growth can now be readily isolated. Furthermore, the entire transfection workflow is greatly accelerated, with recombinant cells that can be generated in days rather than weeks. Another advantage is that molecular genetics can further be performed with freshly isolated wild-type Dictyostelium samples from the environment. This can help to extend the scope of approaches used in these research areas.
The Dictyostelium genus are soil-living social amoeba that mainly feed on bacteria. Being placed in the phylum Amoebozoa, a large number of species have been isolated that can be grouped into four different clades1. The species Dictyostelium discoideum (D. discoideum) has become a popular model organism to study complex cellular processes such as cell migration and phagocytosis. To control and standardize experimental conditions, axenic cell lines have been developed that are able to grow in complex or defined liquid medium in the absence of bacteria2. Of particular importance are the Ax2, Ax3, and Ax4 strains, which were all generated in the 1970s and ultimately derived from a single wild isolate NC43. Tools for genetic engineering were developed in these axenic strains, resulting in the first published knockout in 19874,5. Protocols were further developed and optimized for use under axenic conditions6,7.
Adaptation of these protocols to wild-type D. discoideum strains that are not able to grow in liquid broth have been attempted by several laboratories. However, this has not become fully successful since the transfection protocols are complex and lack efficiency, in part due to the capacity of the bacteria to act as a sink for the selective reagents8,9. As a result, essentially all molecular data on D. discoideum comes from descendants of a single wild-type isolate. We wanted to overcome this limitation and develop a method to genetically modify D. discoideum cells independent of their ability to grow in liquid medium. The need for such a method can be explained by the observation that it was assumed in the past that the mutations allowing axenic growth were mainly neutral and did not impair cell physiology. This supposition is only partially correct. In general, there are two notable differences; first, between the different isolated axenic strains, and second, when these axenic strains are compared with non-axenic wild isolates8,9.
Perhaps the most critical factor is the key axenic gene, axeB, that was identified recently as RasGAP NF1. The major function of NF1 as a RasGAP is to restrain Ras activity3. The deletion of the enzyme in all axenic strains leads to excessive Ras activity manifested as the formation of large active Ras patches. These enlarged Ras patches lead to the accumulation of PIP3 in the plasma membrane. Those coincidental appearing patches of PIP3 and active Ras are a template for the formation of a circular ruffle that eventually closes and leads to the formation of macropinosomes10. The consequence is an excessive increase in macropinocytic activity. Macropinocytosis is an actin-driven process. A competition for cytoskeletal components for the formation of either macropinosomes or pseudopods is the result. Its impact on cell behaviour is reflected in the almost complete prevention of chemotaxis of vegetative cells to folate11. The massively enlarged PIP3 patches are very persistent. Even in starved cells, the PIP3 patches remain and can be misinterpreted as pseudopods, which can cause problems interpreting studies on chemotaxis to cAMP.
In some cases, the NF1 mutation is experimentally useful. This leads us to a second motivation for developing a transfection method for bacterially grown D. discoideum cells, since the increase in macropinocytosis rate makes axenic cells valuable for investigating fundamental aspects of this process12. However, mutation in the genes required for macropinocytosis, such as Ras and PI3-kinases10, have almost abolished axenic growth, making it necessary to manipulate these cells through growth on bacteria. Another reason that renders bacteria-based transfections valuable is the increasing use of Dictyostelids to explore questions in the evolution of multi-cellularity13,14, kin recognition15,16, and altruistic cellular behaviour, which mainly depend on the use of freshly isolated wild-type isolates17. All mentioned research areas can be facilitated by efficient methods for genetic manipulation of wild isolates, which are non-axenic and do not grow in liquid broth.
Our protocols allow for overcoming of the described limitations. Taken together, the possibility of performing genetic manipulations with bacterial grown D. discoideum cells holds benefits for all Dictyostelium researchers, even if it is just the increased speed of the selection process due to faster growth of the amoeba (4 h doubling time) on bacteria compared to growth in axenic media (10 h doubling time).
plasmid name resistance/selection in bacteria resistance/selection in Dictyostelium tag
extrachromosomal expression plasmids
pDM1203 Ampicilin G418 no
pDM1207 Ampicilin G418 N-terminal GFP
pDM1208 Ampicilin G418 N-terminal mCherry
pPI159 Ampicilin G418 N-terminal mNeon
pPI437 Ampicilin G418 N-terminal mScarlet
pPI54 Ampicilin G418 N-terminal mTurquoise2
pDM1209 Ampicilin G418 C-terminal GFP
pDM1210 Ampicilin G418 C-terminal mCherry
pPI143 Ampicilin G418 C-terminal mNeon
pPI459 Ampicilin G418 C-terminal mScarlet
pPI142 Ampicilin G418 C-terminal mTurquoise2
shuttle plasmids
pDM344 Ampicilin no no
pDM1019 Ampicilin no N-terminal GFP
pDM1018 Ampicilin no N-terminal mCherry
pPI152 Ampicilin no N-terminal mNeon
pPI418 Ampicilin no N-terminal mScarlet
pPI150 Ampicilin no N-terminal mTurquoise2
pDM1021 Ampicilin no C-terminal GFP
pDM1020 Ampicilin no C-terminal mCherry
pPI153 Ampicilin no C-terminal mNeon
pPI457 Ampicilin no C-terminal mScarlet
pPI151 Ampicilin no C-terminal mTurquoise2
inducible extrachromosomal expression plasmids
pDM1038 Ampicilin Hygromycin no
pDM1047 Ampicilin Hygromycin N-terminal GFP
pDM1046 Ampicilin Hygromycin N-terminal mCherry
pPI450 Ampicilin Hygromycin N-terminal mNeon
pPI452 Ampicilin Hygromycin N-terminal mScarlet
pPI449 Ampicilin Hygromycin N-terminal mTurquoise2
pDM1049 Ampicilin Hygromycin C-terminal GFP
pDM1048 Ampicilin Hygromycin C-terminal mCherry
pPI470 Ampicilin Hygromycin C-terminal mNeon
pPI460 Ampicilin Hygromycin C-terminal mScarlet
pPI469 Ampicilin Hygromycin C-terminal mTurquoise2
act5 safe haven targeting plasmids
pDM1501 Ampicilin Hygromycin no
pDM1513 Ampicilin Hygromycin N-terminal GFP
pDM1514 Ampicilin Hygromycin N-terminal mCherry
pPI231 Ampicilin Hygromycin N-terminal mNeon
pPI419 Ampicilin Hygromycin N-terminal mScarlet
pPI228 Ampicilin Hygromycin N-terminal mTurquoise2
pDM1515 Ampicilin Hygromycin C-terminal GFP
pDM1516 Ampicilin Hygromycin C-terminal mCherry
pPI230 Ampicilin Hygromycin C-terminal mNeon
pPI458 Ampicilin Hygromycin C-terminal mScarlet
pPI229 Ampicilin Hygromycin C-terminal mTurquoise2
REMI expression plasmids
pDM1220 Ampicilin Hygromycin no
pDM1351 Ampicilin Hygromycin N-terminal GFP
pDM1259 Ampicilin Hygromycin N-terminal mCherry
pPI465 Ampicilin Hygromycin N-terminal mNeon
pPI468 Ampicilin Hygromycin N-terminal mScarlet
pPI466 Ampicilin Hygromycin N-terminal mTurquoise2
pDM1352 Ampicilin Hygromycin C-terminal GFP
pDM1305 Ampicilin Hygromycin C-terminal mCherry
pPI471 Ampicilin Hygromycin C-terminal mNeon
pPI467 Ampicilin Hygromycin C-terminal mScarlet
pPI472 Ampicilin Hygromycin C-terminal mTurquoise2
targeted in frame plasmids
pDM1355 Ampicilin Hygromycin C-terminal GFP
pPI461 Ampicilin Hygromycin C-terminal mCherry
pPI462 Ampicilin Hygromycin C-terminal mNeon
pPI464 Ampicilin Hygromycin C-terminal mScarlet
pPI463 Ampicilin Hygromycin C-terminal mTurquoise2
knock-out plasmids
pDM1079 Ampicilin Blasticidin no
pDM1080 Ampicilin Nourseothricin no
pDM1081 Ampicilin Hygromycin no
pDM1082 Ampicilin G418 no
CRE expression plasmids
pDM1483 Ampicilin Nourseothricin no
pDM1489 Ampicilin Hygromycin no
pDM1488 Ampicilin G418 no
Extrachromosomal plasmids are used for reporter studies, which aim to identify the localization of certain proteins inside a cell or changes in cellular structure of mutant cells. For many approaches, such as monitoring of the cell cycle, it is crucial to express two reporters at the same time. This is now possible using our dual reporter extrachromosomal plasmid system (Table 1). On day 1, cells were transfected before adding the selectable marker G418 after 5 h (Figure 1). In the example, NC4, DdB, Ax2, and the independently derived wild isolates V12M2 and WS2162 (Supplementary Table 1) were transfected with the plasmid pPI289, which encodes for GFP-TubulinA, a marker for microtubules and mCherry-PCNA, a protein that is used for monitoring the cell cycle (Figure 2A). After 32 h, the cells were observed under the microscope. The majority of cells expressed both fluorescent-labeled fusion proteins, consistent with previous reports that expression of two reporters from the same plasmid shows similar expression levels, which is nearly impossible when using two different plasmids18,19. A representative cell for each cell line (NC4, DdB, Ax2, V12M2, and WS2162) expressing the desired dual reporter is shown in Figure 2B. The transfection efficiencies are summarized in Figure2C. NC4-derived cell lines show the best transfection efficiencies. However, for cell lines V12M2 and WS2162, a considerably high number of transfectants was obtained.
Figure 2: Expression of an extrachromosomal plasmid. (A) Extrachromosomal plasmids are directly transfected in circular form. As an example, the dual reporter pPI289 is shown. The NgoMIV sites indicate insertion of the second reporter into the extrachromosomal expression plasmid. (B) Z-projection of a representative cell expressing GFP-TubulinA (cytoplasmic) and mCherry-PCNA (largely nuclear) for five different wild-type cell lines used (NC4, DdB, Ax2, V12M2, WS2162). (C) The transfection efficiencies for the five cell lines pictured in (B) were calculated. Shown is the mean of two experiments. Error bars indicate ± SD. Please click here to view a larger version of this figure.
Integration of a targeting vector into a specified genomic locus is more challenging and requires more careful analysis of the generated cell line. In Figure 3, an act5-mCherry KI in NC4 is attempted. First, the plasmid must be linearized to increase the frequency of recombination events following transfection. For this, the plasmid pDM1514 is cut with NgoMIV. Two bands are obtained after running the digest on an agarose gel. The 4127 bp band contains the desired construct (Figure 3). For transfection, the digested DNA must be extracted from the gel and purified using a gel extraction kit following the manufacturer's instructions.
Figure 3: Preparation of act5 knock-in and DNA for transfection. An example of the use of an act5 knock-in the plasmid pDM1514 is shown. The steps for preparation are listed as follows. (A) Before electroporation, linearize the plasmid using the indicated NgoMIV sites. (B, C) Run the cut plasmid on an agarose gel until the two expected bands are properly separated. Cut out the 4127 bp band containing the recombination arms, mCherry, and the resistance-cassette, and gel extract the DNA. The DNA is now ready for transfection. Please click here to view a larger version of this figure.
The purified DNA was used for transfection of NC4 cells. After 5-6 days of selection, clones were obtained. Representative transfection efficiencies and the amount of positive identified clones for several act5 knock-in attempts are summarized in Table 3. Two clones of the NC4 transfection were randomly chosen and analyzed by PCR (Figure 4A), and both showed the predicted band patterns expected of a knock-in and were further validated with Southern blot analysis to ensure a single integration event of the construct into the genome20. The blot shows a clear single integration in the desired act5 locus (Figure 4B). The generated NC4::act5-mCherry cell line can now be used in experiments.
Figure 4: Validation of act5-mCherry KIs in NC4. (A) Scheme and control PCRs for the validation of positive integrations into act5 locus. The indicated primers were used to analyze two independent clones and the parent. Both clones show the expected bands for the resistance-cassette and downstream (P1) or upstream primer (P2), which are not present in the parental strain. The primer combination (P1/P2) confirms the correct integration of mCherry and resistance cassette into the act5 locus. The wild-type NC4 shows the expected 2800 bp band, while both KI clones lack this band and instead display a PCR product about 1400 base pairs larger. (B) Scheme for the used restriction digest and Southern blot. Both knock-in clones show the smaller 3400 bp band, which is the result of integration of the construct into the act5 locus, specifically the additional BclI site in the hygromycin resistance cassette. The wild-type control shows the expected 5.8 kb resulting from the two downstream located BclI sites. The blot was cropped and spliced for clarity. Please click here to view a larger version of this figure.
Table 3: Transfection efficiencies and the amount of obtained positive transfectants for the generation of act5 KIs in different strain backgrounds22.
The act5 locus offers relatively homogenous expression of the integrated reporter21. The generated NC4::act5-mCherry cells allow mix experiments to be performed with other act5 knock-ins using a different fluorescent protein such as GFP. To emphasize the great advantage of this system, mixing experiments with Ax2::act5-GFP are shown. Due to the inability to transfect non-axenic wild-type cells, this type of approach could not be performed before. Mix experiments are an important tool to analyze cell behavior, because they allow a direct comparison between different cell lines experiencing identical experimental conditions. NC4 cells grow faster on bacterial lawns than Ax2 cells (Figure 5A). This may be due to a higher capacity to phagocytose bacteria or improved ability to move and show chemotaxis towards a food source. Using an under-agar folate chemotaxis assay, direct comparison of a population of NC4::act5-mCherry and Ax2::act5-GFP was performed, showing that NC4::act5-mCherry are much more efficient in folate sensing. After 4 h, more NC4::act5-mCherry cells were able to crawl under the agarose than Ax2::act5-GFP cells (Figure 5B). Analyzing standard metrics of chemotaxis for the cells migrating under the agarose revealed that NC4::act5-mCherry cells were faster and showed stronger chemotactic response than Ax2::act5-GFP cells (Figure 5C-E).
Figure 5: Using act5 KIs for image-based chemotaxis mix experiments. (A) NC4::act5-mCherry and Ax2::act5-GFP expressing the indicated fluorescent protein from the act5 locus were analyzed for the ability to grow on a bacterial lawn. After 4 days, the plaque diameter arisen from solitary plated Dictyostelium cells were measured. Non-axenic act5::NC4 cells make significantly bigger plaques than axenic Ax2::act5-GFP cells (mean ± SD, ***p < 0.0001, n = 3, scale bar 5 mm). (B) For use of the under-agarose folate chemotaxis assay22 to directly compare the chemotactic abilities of the NC4::act5-mCherry and Ax2::act5-GFP used in (A), bacteria- grown amoebae of both strains were mixed in a 50:50 ratio. Cells were allowed to crawl under the agarose. After 4 h, cells that were migrating up the folate gradient were imaged using a confocal microscope. The number of NC4::act5-mCherry and Ax2::act5-GFP cells was then determined. NC4::act5-mCherry cells were more efficient in sensing folate. About 10-fold more NC4::act5-mCherry cells were found compared to Ax2::act5-GFP cells (mean ± SD, ***p < 0.0001, n = 6, scale bar 100 µm). (C, D) The cells were filmed for 60 min, and their speed and chemotactic index were calculated. After pre-selection of the most chemotactically responsive cells (only those that migrated under the agarose), Ax2::act5-GFP cells showed lower values for both cell speed and chemotaxis. Fifty cells per cell line were analyzed. (median ± SD, *p < 0.01, n = 3). (E) The tracks of NC4::act5-mCherry and Ax2::act5-GFPact5 knock-ins are plotted over 60 min, showing the more directed movement towards the chemoattractent source of the NC4::act5-mCherry cells (median ± SD, ***p < 0.0001, n = 6). Please click here to view a larger version of this figure.
The act5 knock-in cell lines can also be used for flow cytometry. As with growth on bacteria, there are major differences in development between axenic strains and non-axenic wild-types. After development, the fruiting bodies of NC4 cells are approximately twice as big as those derived from Ax2 cells. If cells are mixed, an intermediate sized fruiting body is obtained. To analyze the contribution of the both cell lines more quantitatively, flow cytometry was used. These analyses showed clearly that the intermediate-sized fruiting bodies are due to different levels of contribution from both cell lines. While NC4::act5-mCherry cells made up about 75% of the measured spores, Ax2::act5-GFP contributed only 25%, revealing a potential fitness advantage for non-axenic strains (Figure 6). Since the analysis does not monitor the stalk cell population, there are two possibilities to explain the imbalance in development between Ax2 and NC4. One possibility is that Ax2 cells contribute mainly to the stalk cell population, rather than entering the spore cell population. Alternatively, more NC4 cells may enter the developmental cycle, with Ax2 relatively delayed, and they are consequently unable to contribute to the assembly of a fruiting body. The possibility to transfect non-axenic wild-type cells further develops other approaches and greatly simplifies experimental procedures.
Figure 6: Act5 KIs allow analysis of mix experiments using flow cytometry. (A) NC4::act5-mCherry and Ax2::act5-GFP were developed separately or in a 50:50 mixture on non-nutrient agar plates. NC4::act5-mCherry cells form larger fruiting bodies than Ax2::act5-GFP. The mix instead shows an intermediate size (scale bar 5 mm). Confocal fluorescence light microscopy suggests a higher amount of NC4::act5-mCherry spores in the spore heads derived from mixes. (B) To quantify this observation, the amounts of spores in harvested spore heads from the mixing experiment in (A) were analyzed by flow cytometry. About 75% of the spores originated from NC4::act5-mCherry cells, with only 25% from Ax2::act5-GFP cells. (C) Representative distribution of spores from both cell lines shown in a flow cytometry scatter. About 0.05% show positive mCherry and GFP signals, suggesting that parasexual fusion processes have occurred or that spores are sticking to each other. Please click here to view a larger version of this figure.
Supplementary Table 1: Different strains of Dictyostelium studied.
The use of non-axenic, wild-type Dictyostelium cells has been very limited so far in molecular research. Available methods for genetic engineering of these strains have lacked reliability and efficiency23, preventing their general adoption. The generated materials and protocols presented here can be used for any D. discoideum strain independent of its ability to grow in liquid medium. It should be mentioned that this protocol is optimized for NC4-derived cell lines. Transfection efficiencies for freshly isolated strains from the wild differ from NC4, as we have observed before and are shown here for V12M2 and WS21628,9. The electroporation conditions seem in particular to have a considerable influence on transfection efficiencies and may require further optimization for some strains. In general, a sufficient amount of transfectants have been observed in all strains tested so far, showing that these methods are workable. The number of positive transfectants obtained using NC4-derived strains is higher compared to other non-axenic wild-type strains, but in all cases, sufficient numbers of transfectants are obtained to allow for further experiments. This, plus the simplicity of our protocol, is the major improvement compared to earlier attempts8,9.
These new non-axenic protocols cover all standard genetic procedures and add further advantages, since transfections can be performed rapidly and efficiently in parallel. Positive clones can be obtained in days rather than weeks, since growth on bacteria halves the division time. The newly established plasmids also work under axenic conditions and can be routinely used under both growth conditions, which is another advancement of this methology22. As introduced in the protocol, the plasmid used for selection on bacteria have special requirements that are crucial for the success of the transfection. In particular, the promoters driving the expression and resistance cassettes are important for successful transfection. The often used act6 (actin 6) promoter24 for driving the resistance or expression cassettes lacks efficiency when cells are grown on bacteria. In our plasmid system, the highly active act15 (actin 15) promoter drives all expression cassettes, while the resistance cassettes are under the control of a coA (coactosin A) promoter, both of which are active under axenic conditions and in cells grown on bacteria. Requirements for the expression of reporter constructs as well as knock-in and knock-out constructs make it necessary to use our plasmid repertoire, but unfortunately limits the use of vectors already created that use the inefficient act6 promoter.
Promoter efficiencies are especially critical for transfections that depend on a single integration event into the correct genomic locus. Due to our improvement of promoters driving the resistance gene expression, enough resistance protein is produced from single locus integrations. A major concern was the favoring of multiple integrations into the genome when using hygromycin or G418 as described. No favoring of multiple integrations has been observed so far, as reported previously for G418 selections using older promoter systems25. This means that both hygromycin and G418 are suitable selectable markers for the generation of clean knock-outs and knock-ins. Unfortunately, the selectable marker blasticidin does not work under non-axenic conditions. This is a major disadvantage of our method, since the blasticidin resistance cassette is routinely used to generate knock-out constructs in D. discoideum. Constructs that were already generated with blasticidin resistance cassettes will need to be rederived using one of the workable selectable markers. Another possibility to overcome this limitation is to combine the recently established axenic D. discoideum CRISPR technology with this transfection protocol26. The generation of suitable, single guide RNAs (sgRNAs) is simpler and faster than the reconstruction of complete knock-out constructs. For future directions, the possibility of generating multiple knock-outs using CRISPR/Cas9 combined with this transfection protocol is appealing and may pave the way for many researchers in the Dictyostelium community. However, the workability of the established transient expression system used for CRISPR/Cas9 in axenic grown cells should be carefully examined.
The act5 knock-in system presented offers a reliable and safe integration system for the generation of stable cell lines in D. discoideum, with the advantage of similar sites established in other organisms22. It also depends on a single integration event in the genome and offers many possibilities for different research directions. The act5 promoter is strongly active and guarantees almost homogenous expression27 independent of the cultivation conditions. Fluorescent reporter proteins can easily be integrated into this safe haven locus using the engineered targeting plasmids. This can be useful, for example, for cell-tracking purposes, as shown here in a mix experiment. The cells show minimal cell-to-cell expression variability, which can assist in automated cell tracking. Importantly, the insertion of a desired sequence into the act5 locus appears phenotypically neutral28,29. As the expression does not depend on a selectable marker to maintain protein expression, as seen in random integrants or extrachromosomal vector-bearing cell lines, the act5 system can be useful for rescue experiments, as well. Since the cell lines are homogenous, all cells can be analyzed. This is not possible using an extrachromosomal plasmid for expression, in which there is considerable heterogeneity in expression.
In addition to these general improvements for all strains, the ability to use non-axenic wild-type cells for molecular research allows an assessment of the effects of accumulated mutations in current laboratory strains. Multiple genetic rearrangements has been found since the adoption of the Ax2 strain in 1970, as shown by previously published microarray analyses26. Synthetic phenotypes are observed because of the presence of these mutations. For example, a reported phg2 mutant shows decreased adhesion in one laboratory strain background and increased adhesion in another3. Such conflicting data can now be resolved by repeating the experiment in the common ancestor strain (in this case, DdB). The strength of this approach has been recently shown for the small GTPase RasS30,31.
The ability to perform genetic experiments in any D. discoideum strain expands the potential of new research directions that depend on the use of wild isolates, notably those involving social evolution, kin recognition, and the sexual cycle22.

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