Non-model model organisms

Abstract

Model organisms are widely used in research as accessible and convenient systems to study a particular area or question in biology. Traditionally only a handful of organisms have been widely studied, but modern research tools are enabling researchers to extend the set of model organisms to include less-studied and more unusual systems. This Forum highlights a range of 'non-model model organisms' as emerging systems for tackling questions across the whole spectrum of biology (and beyond), the opportunities and challenges, and the outlook for the future.

Fig. 1

Images of various diatom species. a Differential interference contrast images of (clockwise from top left): Striatella unipunctata, Odontella sp., Stephanopyxis turris, Pseudo-nitzschia sp., Thalassiosira sp., Cylindrotheca sp., Asterionellopsis glacialis, Skeletonema costatum, Grammatophora oceanica, and Chaetoceros sp. Images are courtesy of Colleen Durkin and reproduced from [324]. b Differential interference contrast image of Coscinodiscus excavatus, image courtesy of Robert Lavigne. c–d Scanning electron micrographs of Stephanopyxis turris theca (c) and nanoscale features (d), images courtesy of Mark Webber. e-f Cylindrotheca fusiformis before cell division (e) and during cell division (f). Top: phase contrast. Bottom: polymerized silica labeled with HCK-123 dye (green) and endogenous chlorophyll fluorescence (red). Scale bar in b 20 μm

Fig. 2

Single-cell regeneration in Stentor coeruleus a A living Stentor cell. The oral apparatus, located towards the upper left of the image, is a large ring of cilia that collects food particles from the surrounding pond water. At the other end of the cell, a holdfast attaches the cell to the surface of pond plants. b Regeneration after bisection of a Stentor cell. The panel on the left shows the longitudinal strips of blue pigment that serve as markers for cellular pattern. When a Stentor cell is cut in half with a glass needle, as indicated by the dotted orange line, each half initially heals its wounds to prevent cytoplasm from leaking out (middle panels) and then within approximately one day, regenerates a complete cell (right panel), with the anterior half regenerating a new holdfast, and the poster half regenerating a new oral apparatus. Both halves are able to regenerate because the cell contains a long polyploid macronucleus running down the length of the cell, such that when a cell is cut, both halves retain many copies of the genome

Fig. 3

Oxytricha as a model system to study genome biology, epigenetic inheritance, and somatic differentiation. a-c Single (a,c) and mating (b) Oxytricha cells. Blue indicates DNA; Yellow is tubulin, highlighting the cilia. i  =  micronucleus, a  =  macronucleus. Image in (a) courtesy of National Human Genome Research Institute, (b) courtesy of Robert Hammersmith, Ball State University and (c) courtesy of Wenwen Fang and reproduced from [325]. d The sexual life cycle of Oxytricha trifallax and rearrangement of scrambled genes, reproduced from [47]. All vegetative cells (stages 1 and 10) contain one (bi-lobed) macronucleus (MAC) and two micronuclei (MIC). The two MIC are genetically identical, but for simplicity we show only one here. (2) When starved, two cells of compatible mating types partially fuse to initiate conjugation. Cell fusion occurs soon after mixing of mating types. (3) Both vegetative micronuclei in each partner enter meiosis I. (4) One product from each meiosis I is promoted to meiosis II, and one of the four meiosis II products is promoted to a post-meiotic mitosis. (5) The sister products of one mitosis develop into gametic nuclei: a posterior stationary and an anterior migratory nucleus. This happens in both partners, such that both cells emerge with identical zygotic genotypes after the exchanged migratory nucleus fuses with the retained stationary nucleus (6), resulting in (7) two genetically identical exconjugant cells. (8) The newly formed zygotic nucleus divides twice: one daughter nucleus is destroyed, two become the new micronuclei, and (9) the parental macronucleus in each partner cell degrades, leaving telomere-to-telomere RNA transcripts behind to guide rearrangement [44, 45]. One zygotic nucleus differentiates into the new macronucleus. This cycle takes approximately 48–60 h. Shown inside the circle are representative MIC and MAC versions of a scrambled gene. Retained DNA segments in purple; deleted DNA regions, including flanking DNA, in yellow; numbered segments correspond to the order in the expressed MAC version; segment 2 is inverted; telomeres are shown as black bars at the ends of the MAC chromosome

Fig. 4

Naegleria gruberi cells undergo a dramatic transformation between crawling amoebae and swimming flagellates, assembling an entire microtubule cytoskeleton along the way. The crawling amoebae (top left) lack cytoplasmic microtubules, but use their actin cytoskeleton (pink) to crawl with two types of protrusions (insets): actin-filled pseudopods and cytoplasm-filled spheres called blebs that appear after delamination of the cell membrane from the underlying actin cortex. Amoebae can respond to a variety of environmental signals by differentiating into a vigorously swimming flagellate (upper right). This process requires the transcription, translation, and assembly of an entire microtubule cytoskeleton (green), including tubulin. Amoebae also can undergo a closed mitosis (lower left), during which the nuclear envelope remains intact, isolating the spindle microtubules (blue) from the cytoplasm. Mitotic microtubules are thought to be built from divergent tubulin isoforms that are expressed prior to mitosis, and then rapidly degraded after cytokinesis

Fig. 5

R-bodies transition between two states. a Type 51 R bodies reversibly transition between a rolled state at neutral pH and a tube state at low pH by the extension of the ribbon from the center of the coil. The state depicted in the middle is transient. b This transition is visible macroscopically; coiled R bodies sediment and extended R bodies remain in solution. c When an R-body-containing bacterium is ingested by a sensitive strain of paramecium after being shed by a “killer” strain, the extension of the R body is triggered by the acidic environment of the food vacuole. Extension causes the rupture of the bacterium and the food vacuole, releasing co-expressed toxins that ultimately result in the death of the sensitive paramecium. Images adapted from [89]

Fig. 6

Schizosaccharomyces japonicus and Schizosaccharomyces pombe exhibit divergent mitotic programs. Left: Live S. pombe and S. japonicus cells expressing the endoplasmic reticulum marker GFP-ADEL and the nucleoplasmic protein Nhp6-mCherry. Note a considerably larger cell size in S. japonicus. Scale bar 5 μm. Right: Schematic representation of mitotic division in the two sister species. Adapted from [108], Current Opinion in Microbiology, Vol 28, Gu, Y. and Oliferenko, S., Comparative biology of cell division in the fission yeast clade, p.18-25, Copyright (2015), with permission from Elsevier

Fig. 7

Ashbya gossypii as a model for cytosolic organization. Left: image of a growing young mycelium. Middle: A.gossypii hyphae with clustered mRNA transcripts. Asynchronously cycling nuclei are shown in blue and clustered cyclin transcripts in orange. Right: cartoon depiction of A.gossypii hyphae with nuclei and clustered transcripts. Scale bars 5 μm

Fig. 8

Volvox and volvocine algae. a Cladogram of selected volvocine species shown in cartoon form with successive cellular and developmental innovations indicated by bulleted descriptions above or below the node in which they arose. Species with published sequenced genomes have names in blue-shaded boxes. b–e Light micrographs of vegetative phase Volvox carteri (Volvox) showing a mature pre-cleavage stage adult (b); a mother spheroid with juveniles (c); and an isolated gonidium (d) or somatic cell (e) from a mature pre-cleavage adult spheroid. f Light micrograph of a Chlamydomonas reinhardtii cell. g Schematic of the Volvox vegetative life cycle synchronized to a 48-h diurnal cycle. A boxed key showing cell types and extracellular matrix (ECM) is in the upper left. Development starts with mature pre-cleavage adults (~11:00 on diagram) and proceeds clock-wise through embryogenesis, cyto-differentiation of germ cells (gonidia) and somatic cells in juveniles, hatching of juveniles, and maturation to become the next generation of adults. After hatching the ECM and parental somatic cells of the previous generation are discarded. The cartooned stages corresponding to light micrographs in panels b and c are labeled

Fig. 9

Haploid tissues of the moss Physcomitrella patens. A plant regenerated from protoplasts is shown in the top center. The boxed regions in this image represent the juvenile (protonemata) and adult (gametophore) tissues, which are drawn schematically on either side of the image. Images acquired from tissue grown in microfluidic devices showing a variety of cell types and tissues are shown in the bottom row

Fig. 10

Cerebral organoids model the architecture of the developing human brain. Left: a section of an entire cerebral organoid stained for the forebrain marker Foxg1, the intermediate progenitor marker Tbr2, and DAPI, revealing the presence of lobules of cerebral cortex as well as other brain regions not positive for Foxg1. Right: a schematic of a lobule of cortex in an organoid showing the proper organization of progenitor zones: ventricular zone (VZ) where radial glial neural stem cells reside, subventricular zone (SVZ) where transit amplifying populations reside, and the intermediate zone (IZ) and cortical plate (CP) where neurons migrate to their final positions. Scattered pink puncta represent outer radial glia, a population abundant in human brain development but much less present in rodents, while elongated purple neurons represent tangentially migrating interneurons that originate outside the cortex. In the case of microcephaly (lower left) organoids overall are much smaller, as are progenitor zones [192], whereas organoids derived from autistic patients display increased numbers of interneurons [198]. Scale bar 100 μm

Fig. 11

Nematostella vectensis phylogenetic position and juvenile morphology. a Metazoan phylogenic tree highlighting the position of Nematostella. The detailed ingroup relationships of medusozoa, as well as the position of Ctenophora and Porifera, are still uncertain, as indicated by question marks. b Morphology of a fully relaxed juvenile Nematostella vectensis polyp. The thickened internal foldings along the body column are called mesenteries. These delicate structures contain digestive glands, and retractor muscles as well as gonads. The reddish coloration of mesenteries is due to feeding of Artemia nauplii under laboratory conditions. Tree in a adapted from [209, 326, 327]: Bridge, D; Cunningham, C W, Class-level relationships in the phylum Cnidaria: molecular and morphological evidence, Molecular Biology and Evolution, 1995, Volume 12, Issue 4, p.679-89, by permission of Oxford University Press and Society for Molecular Biology and Evolution

Fig. 12

Scanning electron micrograph of the water bear Hypsibius dujardini. Image credit: Vicky Madden and Bob Goldstein

Fig. 13

Limb regeneration in salamanders. Reprinted from [328], Elsevier Books, Richard Goss, Principles of Regeneration, Copyright (1969), with permission from Elsevier

Fig. 14

The African turquoise killifish has two distinct phases in its lifecycle. During the rainy season, the turquoise killifish has a naturally compressed lifecycle. Turquoise killifish grow fast and rapidly reach sexual maturation, characterized by bright colors in males (Young adult). Old fish recapitulate aging phenotypes, including loss of muscle mass, color, and tissue homeostasis (Old adult). Newly laid turquoise killifish embryos enter diapause to survive the upcoming drought during the dry season (embryo). The embryos can stay in diapause for many months (several times longer than the fish lifespan), raising the possibility that the damage that accumulates with time may be slowed or even reset (“rejuvenation”). The embryos then break diapause and the fish resume their compressed lifecycle during the following rainy season. Some embryos escape diapause, and it is therefore possible to study each state separately in the laboratory