Roles Conceptualization, Data curation, Formal analysis, Methodology, Validation, Visualization, Writing – original draft, Writing – review & editing * E-mail: a.alcalde@bristol.ac.uk (AAA); s.montgomery@bristol.ac.uk (SHM) Affiliation School of Biological Sciences, University of Bristol, Bristol, United Kingdom
Contributed equally to this work with: Max S. Farnworth, Laura Hebberecht Roles Methodology, Writing – review & editing Affiliation School of Biological Sciences, University of Bristol, Bristol, United Kingdom
Contributed equally to this work with: Max S. Farnworth, Laura Hebberecht Roles Methodology, Writing – review & editing Affiliation School of Biological Sciences, University of Bristol, Bristol, United Kingdom ⨯
Roles Conceptualization, Supervision, Writing – review & editing Affiliation School of Biological Sciences, University of Bristol, Bristol, United Kingdom ⨯
Roles Conceptualization, Funding acquisition, Investigation, Supervision, Writing – review & editing * E-mail: a.alcalde@bristol.ac.uk (AAA); s.montgomery@bristol.ac.uk (SHM) Affiliation School of Biological Sciences, University of Bristol, Bristol, United Kingdom
29 Aug 2024: The PLOS ONE Staff (2024) Correction: A modified method to analyse cell proliferation using EdU labelling in large insect brains. PLOS ONE 19(8): e0309825. https://doi.org/10.1371/journal.pone.0309825 View correction
The study of neurogenesis is critical to understanding of the evolution of nervous systems. Within invertebrates, this process has been extensively studied in Drosophila melanogaster, which is the predominant model thanks to the availability of advanced genetic tools. However, insect nervous systems are extremely diverse, and by studying a range of taxa we can gain additional information about how nervous systems and their development evolve. One example of the high diversity of insect nervous system diversity is provided by the mushroom bodies. Mushroom bodies have critical roles in learning and memory and vary dramatically across species in relative size and the type(s) of sensory information they process. Heliconiini butterflies provide a useful snapshot of this diversity within a closely related clade. Within Heliconiini, the genus Heliconius contains species where mushroom bodies are 3–4 times larger than other closely related genera, relative to the rest of the brain. This variation in size is largely explained by increases in the number of Kenyon cells, the intrinsic neurons which form the mushroom body. Hence, variation in mushroom body size is the product of changes in cell proliferation during Kenyon cell neurogenesis. Studying this variation requires adapting labelling techniques for use in less commonly studied organisms, as methods developed for common laboratory insects often do not work. Here, we present a modified protocol for EdU staining to examine neurogenesis in large-brained insects, using Heliconiini butterflies as our primary case, but also demonstrating applicability to cockroaches, another large-brained insect.
Citation: Anton AA, Farnworth MS, Hebberecht L, Harrison CJ, Montgomery SH (2023) A modified method to analyse cell proliferation using EdU labelling in large insect brains. PLoS ONE 18(10): e0292009. https://doi.org/10.1371/journal.pone.0292009
Editor: Gregg Roman, University of Mississippi, UNITED STATES
Received: April 5, 2023; Accepted: September 11, 2023; Published: October 5, 2023
Copyright: © 2023 Anton et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: This is a lab protocol which does not report data.
Funding: This work was funded by an ERC Starter Grant (758508) and a NERC IRF (NE/N014936/1) to SHM. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
The study of neurogenesis is critical to understanding the evolution and development of nervous systems. Neurogenesis is the process by which neural progenitor cells (neuroblasts) divide and ultimately generate neurons, a process with common features across vertebrates and invertebrates [1]. In invertebrates, this process has been most intensely studied in the fruit fly, Drosophila melanogaster [2–5], owing to the exhaustive availability of genetic tools, but there are also isolated but key insights provided by other insect species that reveal conserved and divergent features of brain development [6–8]. For example, consistently across insect species [7, 9–12], so-called type I neuroblasts (NB), divide asymmetrically multiple times, generating one ganglion mother cell (GMC) with each division, and self-renewing for the next cycle. GMCs subsequently produce two identical cells, either neurons or glial cells. Adult cell number is therefore largely determined by the number of rounds of neuroblast division. Type II neuroblasts, which account for a relatively small number of cell lineages in Drosophila [13], instead divide symmetrically to generate another cell type, intermediate progenitor cells (IPCs). These IPCs subsequently divide asymmetrically leading to self-renewal and a GMC, which terminally divides symmetrically producing two neurons. Adult cell numbers in these lineages are therefore determined by the number of rounds of neuroblast and IPC division. Hence, by adding a second proliferative phase IPCs increase the final number of neurons produced by neuroblasts [14–17].
Nervous systems in insects are extremely diverse in their size, structure and ontogenetic trajectories [18, 19]. This variation is likely explained by altered dynamics of cell proliferation, including variation in neuroblast number, the length of neurogenic cell division, and the propensity to produce IPCs. For instance, D. melanogaster has 4 mushroom body neuroblasts per hemisphere whereas the honey bee, Apis mellifera reportedly has 500 mushroom body neuroblasts. Another axis of interspecific variation can be seen in the timing of key neurodevelopmental events [20]. This diversity of insect neurodevelopment offers the possibility of uncovering mechanisms that govern key cellular processes, leading to a better understanding of the development and evolution of nervous systems. However, to exploit this diversity we must overcome the challenge of developing or optimising methodologies for less frequently studied organisms, for which protocols developed in more common lab insects may be ineffective.
One example of insect nervous system diversity is the size of a central brain structure called the mushroom bodies. Mushroom bodies are one of the most prominent and variable structures in the insect brain. They have a variety of functions, including sensory integration, filtering and attention [21–23]. However, they are particularly implicated in learning, memory, and generally complex behaviours that require the integration of innate states and sensory stimuli [19]. Across insects, mushroom bodies have changed in size multiple times, with particularly large mushroom bodies evolving in at least four lineages [24–27] providing opportunities to study the convergent evolution of expanded neural structures. This size variation reflects changes in the number of intrinsic mushroom body neurons, the Kenyon cells [24, 28]. The developmental mechanisms behind these different population sizes remain largely unclear. Studying the rate and duration of neurogenesis in a comparative context across related species with diverse neural morphologies could advance our understanding of the developmental mechanisms controlling cell production.
To explore variation in Kenyon cell production we focus on one of the four noted increases of mushroom body size, which occur in passion vine butterflies, Heliconius [26, 29]. Relative to the rest of the brain, Heliconius mushroom bodies are 3-4X larger than other genera in the tribe Heliconiini [26, 28, 29]. The close relatedness of the Heliconiini [30], and the general similarity in their ecology [31] and juvenile life history [32], provides a clear opportunity for comparative studies of development. However, this system lacks basic tools to study neurogenesis.
In other insects, three methods have been used to identifying neurogenesis: i) methods based on the incorporation of chemical markers during the S phase or M phase of the cell cycle; ii) methods that use markers against specific proteins expressed in the membrane of proliferating cells; and iiii) genetic tools. The last two groups of methods have primarily been developed for Drosophila, with genetic tools currently being less tractable in other systems. We therefore focused on the first group of methods which can also provide information about cell activity.
The first method used to detect proliferating cells in this way was [ 3 H]-thymidine autoradiography [33]. In this technique [ 3 H]-thymidine binds to the DNA of cells undergoing mitosis and labelled DNA is detected by autoradiography [33]. With this method, Altman reported adult neurogenesis in the human dentate gyrus [34]. The main limitations of this technique are the requirement for use of a radioisotope and the time that the detection takes, which can last months [35]. A related method to label cells in mitosis uses 5-Bromo-2-deoxyuridine (BrdU), a synthetic analogue of thymidine, that binds to DNA during the S phase. This method is faster and provides higher temporal and spatial resolution. It has been widely used to mark neuroblasts in the insect brain [36–39] and to study adult neurogenesis in both vertebrates [40] and invertebrates [37, 41], including moths [42]. Apart from being technically difficult, the main limitation, in this case, is that it requires a strong denaturization of the DNA which can degrade the structure of the sample and adversely affect the tissue’s morphology.
More recently a similar method has been developed that uses an alternative nucleotide analogue, 5-ethynyl-2-deoxyuridine (EdU) [35]. EdU is similar to BrdU but the nucleotide is detected by a chemical reaction. Compared with BrdU this method is more sensitive, faster and does not require DNA denaturalization which permits better conservation of cellular structure [35]. Additionally, studies comparing EdU and BrdU have indicated that EdU is more effective at detecting cell proliferation and easier to use [43, 44]. Other markers such as the anti-phospho-Histone H3 (Ser10) antibody have been used to mark cells in mitosis. The process of phosphorylation of Histone H3 starts during the G2 phase but gets reversed at the end of mitosis. Therefore, this label serves as a momentary "snapshot" rather than a long-lasting indicator. In comparison with pH3, EdU marks every cell undergoing S-phase, so it captures a wider picture of divisions instead of smaller windows of the cell cycle. It has also been used to study neurogenesis [7], but currently in a restricted range of small insect species which are commonly used as model organisms, Drosophila melanogaster and the red flour beetle, Tribolium castaneum [7, 45]. To our knowledge, in Lepidoptera and other large insects, only [ 3 H]-thymidine, BrdU, and histological images have been used to identify dividing cells and their progeny [42, 46, 47]. Although these methods have been very useful, they are limited by the time required, lower sensitivity and, in the case of the histological images, the lack of information about cell activity. In this work, we therefore adapt existing EdU staining protocols to study neurogenesis in large-brained insects, using Heliconiini butterflies as a focal case study.
The protocol described in this peer-reviewed article is published on protocols.io (https://dx.doi.org/10.17504/protocols.io.n92ldmy69l5b/v1) and is included for printing purposes as S1 File.” Experimental variations in the protocol and their outcome are summarized in S1 Table in S2 File and EdU incorporation is further explained in this section. S1 Table in S2 File shows the different combinations trialled here, including those which were successful and unsuccessful, to increase efficiency when designing future experiments.
As a positive control in early experiments we used Oregon R. wild-type flies of Drosophila melanogaster (2,500 Kenyon cells/hemisphere [48]) kept in standard laboratory conditions at 25ºC. We selected and collected flies in the prepupal stage for our experiments. We included comparisons between two Heliconiini, the red postman butterfly, Heliconius erato (52,000 Kenyon cells/hemisphere [48]) and the flame butterfly, Dryas iulia (13,000 Kenyon cells/hemisphere [48]) obtained from breeding stocks established from commercial pupae suppliers (The Entomologist Ltd, East Sussex, UK). Butterflies were maintained in ~2m x 2m x 2m cages at 24ºC– 30ºC and 80% humidity 80%. Each cage contained natural host plants for each species, Passiflora biflora and P. triloba for D. iulia, and P. biflora for H. erato. Butterflies were fed every other day with a pollen/sugar solution (5% pollen or artificial amino acids source, 20% sugar, 75% water). Fresh flowers were also provided from Lantana and Psiguria as additional sources of food. Larvae were reared in individual pots and fed every day with fresh leaves. Young pupae (0–1 days old) were collected to test EdU staining protocols. To provide an additional comparison of whether the protocol developed for Heliconiini worked in other large-brained insects, we obtained 2–4 weeks old Pacific beetle cockroach, Diploptera punctata. Cockroaches came from stock populations at the University of Bristol maintained at 26 ºC and 60% humidity.
The protocol is a modified version from the instructions of the Click-iT ™ EdU Cell Proliferation Kit for Imaging, Alexa Fluor ™ 488 dye (Thermofisher, #C10420). The first step of EdU staining protocols is the incorporation of the EdU nucleotide into the DNA of replicating cells. It therefore requires the cells to be alive for the duration of the incubation. Based on previous studies in Drosophila and Tribolium [7, 45] we diluted EdU in 0.1M PBS in three different concentrations: 10 μM, 20 μM and 50 μM. Pilot experiments in Drosophila led to a final dilution of 20 μM for subsequent trials. As an alternative medium, we tested the benefits of replacing the dilution buffer with Grace’s Medium (ThermoFisher, #11595030) which is used to maintain insect cell cultures [49].
To refine the EdU protocol we focused on larvae and early pupae, where we anticipated high rates of neurogenesis. We tested four main ways of incorporating EdU (Fig 1).