Information

Effect on gene loss because of compartmentalisation of plastids/mitochondria/endosymbiont?


Considering the transfer of genes during endosymbiosis a gene transfer event (at least fundamentally, even if it's a special case), how does the fact that in this case the genes are inside a compartment, affect their gene loss, or incorporation into the host/recipient's genome (in context of gene transfer)?

I would imagine that the loss would be slower, recombination with other transferred genes less probable. But I can't find literature on the same. Any leads?


Integration of plastids with their hosts: Lessons learned from dinoflagellates

After their endosymbiotic acquisition, plastids become intimately connected with the biology of their host. For example, genes essential for plastid function may be relocated from the genomes of plastids to the host nucleus, and pathways may evolve within the host to support the plastid. In this review, we consider the different degrees of integration observed in dinoflagellates and their associated plastids, which have been acquired through multiple different endosymbiotic events. Most dinoflagellate species possess plastids that contain the pigment peridinin and show extreme reduction and integration with the host biology. In some species, these plastids have been replaced through serial endosymbiosis with plastids derived from a different phylogenetic derivation, of which some have become intimately connected with the biology of the host whereas others have not. We discuss in particular the evolution of the fucoxanthin-containing dinoflagellates, which have adapted pathways retained from the ancestral peridinin plastid symbiosis for transcript processing in their current, serially acquired plastids. Finally, we consider why such a diversity of different degrees of integration between host and plastid is observed in different dinoflagellates and how dinoflagellates may thus inform our broader understanding of plastid evolution and function.

Plastids evolve through the endosymbiotic integration of two organisms: a eukaryotic host and a photosynthetic prokaryotic or eukaryotic symbiont. It is generally believed that the host initially consumes the symbiont through phagocytosis. Subsequently, over long evolutionary timescales, pathways evolve within the host to maintain the endosymbiont as a permanent, intracellular organelle (1). At least eight distinct plastid endosymbioses have been documented across the eukaryotes, giving rise to a diverse array of different photosynthetic lineages (reviewed in ref. 2). Understanding what processes underpin the integration of plastids with their hosts may provide valuable insights into the evolution and function of photosynthetic eukaryotes.


Access options

Get full journal access for 1 year

All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.

Get time limited or full article access on ReadCube.

All prices are NET prices.


Mechanisms facilitating endosymbiosis

Mutualism often originates from asymmetrical, even exploitative interactions [12] most of them are facultative, and many have relatively recent origins [65]. Obligate mutualisms are rare and considered less stable, since there is a higher chance of (functional) degradation by occasional loss of partner [50, 65, 70]. Symbiosis is shaped by conflicts of interests which are probably harder to manage at the early stage of the association [14, 71]. Consequently, it is unlikely that the ancestors of mitochondria and host first met with perfect metabolic complementarity, so that their symbiosis was immediately mutually beneficial. On the other hand, these specialized obligate symbioses do exist and are persistent for millions of years despite any conflicts, indicative of stabilizing mechanism. In turn, we will discuss mechanisms that can stabilize an emerging though suboptimal interaction so that in can be selected for.

Group selection in endosymbioses

If groups (associations) form among cells and these groups affect the selection of individual cells, selection appears at multiple levels: individual selection favors the interest of individual cells, while group selection acts in the interest of the associations [11, 72, 73], e.g., of symbiotic pairs. However, multilevel selection almost inevitably leads to between-level conflicts. To better understand group formation, multilevel selection was conceptually characterized into two types, multilevel selection 1 and 2 (MLS1, 2).

In case of MLS1, only temporary groups form that periodically disappear to revert to an unstructured population of cells (also called transient compartmentation) [74]. Facultative (endo- or ecto-) symbioses realize MLS1: partners re-associate better-than-random. Fixed spatial structure of cells can also act as implicit group structure. In dense biofilms, cells are practically immobile and the limited diffusion of exchanged molecules localizes interactions. As a result, mutualist partners stay close and their implicit group can withstand cheating mutants or harmful competitors appearing at group edges [75]. In this case, splitting up the population into explicit, reproductively isolated groups is not required for selection to prefer mutualists [76]. It is yet unknown if endosymbiosis could or have ever evolved in biofilms.

MLS2, on the other hand, involves explicit group structure, i.e., groups that last and reproduce indefinitely. In symbiosis terms, this means exclusive partnership with strict vertical inheritance. If the group is selected for and can stably inherit group-related adaptations, it is a bona fide evolutionary unit (an informational replicator [77]). When obligate codependence of endosymbiotic partners is established, a new unit of evolution emerges [78] and selection of associations dominates over selection of individuals. A major evolutionary transition happens when multilevel selection results in irreversible coupling where individuals forfeit their autonomous replication and gives rise to an association with potential for higher complexity [11]. For group selection to be effective, group members must reproduce together better than random and there must be a selective advantage at the group level. In turn, we will discuss mechanisms that can ensure positive assortativity of partners.

The theory of group selection predicts that the group is favored by selection over individuals if there is a reasonable selective advantage for the group, even if the net of benefits and costs is negative at certain times (i.e., the per capita growth rate of the association is smaller than that of individual cells under certain conditions). Accordingly, the initial partnership does not need to be directly mutually beneficial for both parties at all times, as long as the partnership together enjoys selective advantage averaged over some time or over different environments. Nevertheless, there must be at least a hidden benefit for each party, so that reduced mean fitness in certain periods is compensated. There are at least two general mechanisms to draw such indirect benefits. One is to exploit heterogenous environments, for example temporally fluctuating or spatially differentiated, so that the mean fitness over a wider temporal or spatial range is larger than those of competitors. The other is bet-hedging, that compensates a reduced mean fitness with reduced fitness variance, e.g., with wider tolerance of harsh conditions [79]. This renders the species less prone to extinction in certain selective environments that are truncating, though rare. A prudent strategy counters or even anticipates the effects of a heterogenous environment (see the farming hypothesis, explored theoretically [53]).

Partner choice mechanisms

Pre- or post-infection partner choice can stabilize (partially) beneficial interactions [80]. Pre-infection partner choice is based on cues or signals or screening mechanisms to filter partner quality before actually establishing any association with the partner [65, 80, 81]. Quorum sensing, including intra- and inter-species communication, exists both in bacteria and archaea [82, 83]. There is, however, no guarantee that interacting cells are indeed of the cooperative type, as cheating in the form of dishonest signals can arise [84, 85]. Signals can be of two types: diffusive or contact molecules. Surface contact requires close proximity and these signals are usually partner-specific. Diffusive signal molecules can reach a larger number of cells, but are less effective (being diluted easily) and are usually not partner-specific, hence are less reliable. The specificity and reliability required for obligate pairwise symbiosis suggest that surface contact is preferred over diffusive signals (Fig. 4).

Basic steps of endosymbiosis and organellogenesis. Geometric shapes represent various benefits (e.g., metabolites), solid black arrows represent the source and flow of the various benefits, dashed arrows indicate investments, and colored arrows indicate the option to leave the host. Note that the last step, if involves nuclear integration and protein import, is irreversible

Post-infection partner choice is based on conditional investments, and involves various rewarding or sanctioning mechanisms, including the selective termination of the interaction and the possibility of switching partners [65, 86]. The prerequisite of post-infection partner choice is spatial separation of the multiple partners, so that the host can differentiate and then selectively treat high- or low-quality partners a set-up often referred to as biological markets [13, 80, 87, 88]. The quality of preferable partners depends on multiple factors [65, 87, 88], and often, a low-quality partner is better than no partner at all.

For most of the cases, there is an asymmetry between the mutualist partners in many aspects, such as power of control over the partner, strategic options, availability of alternative partners, etc.[86]. The party with more power or control is expected to gain the higher profit from the interaction, which can even drive the interaction towards unilateral exploitation [58, 65, 88]. Nevertheless, such selectivity by partner control mechanisms can shift the balance in favor of high-quality partners in the population in spite of their competitive inferiority to low-quality partners without the intervention of the mutualist [65]. Additionally, control mechanisms may allow the host to manipulate symbiont behavior and to force higher returns from investing into the symbiont, and may also allow for context-dependent treatment of the partner [56, 58, 89].

Partner fidelity feedback and internalization

Partner fidelity mechanisms are able to reduce the conflict of interest between partners as the symbiont survival depends on the survival of the host [90]. Increasing investment toward the partner increases the amount or possibility of reciprocated investment, i.e., it is a favor returned [48, 80]. The higher the quality of the mutualist, the higher the chances for survival [65]. Such feedbacks can be interpreted in two time-frames: in-generation or cross-generation. In a generation of a long-term partnership, increasing investments induce higher rates of nutrient flows (in nutrition mutualisms) or higher quality of services (in protection mutualisms) by the partner. Partner fidelity can also manifest as a cross-generational effect, where the investment into a high-quality partner will also benefit the progeny [65].

Cross-generational partner fidelity is usually coupled with vertical (or pseudo-vertical) transmission mechanisms, and is similar in effect to spatial structure: it ensures that offspring can form associations with the same selection of partners as parents did. Strict vertical transmission is very rare (besides endosymbiotic organelles, and some cases of parasitism, like Wolbachia in wasps [48, 91]). Imperfect correlation between partners across generations, called pseudo-vertical transmission, is more frequent [48, 92]. Such loose correlations and feedbacks can stabilize mutualism and pave the way for the evolution of perfect cross-generational correlation of partners.

Theory predicts that the evolution of symbiont capture and vertical transmission is driven by host mechanisms to control symbiont transmission [93]. First, because symbiont capture involves the genome reduction of the symbiont while providing increasingly more benefit to hosts, second, because the processes during cell division affecting the distribution and the frequency of reproduction of both parties are controlled by the host, which thus can have the power of selecting which symbionts to transfer (probably restricted to multicellular eukaryotes, e.g., in Buchnera–aphid interactions [94].

Undoubtedly, physical inclusion is the most advanced method of vertical transmission, but at the start of a symbiotic partnership, it is rarely available. In most prokaryotic symbioses, physical inclusion never happens, or is limited to a periplasmic space (e.g., Bdellovibrio [95], Chlorochromatium aggregatum [96]). There are some rare cases where the symbiont can enter the host’s cytoplasm, but, e.g., parasitic Daptobacter ultimately kills its host [97]. Phagotrophic eukaryotes could store their captured symbionts in phagosomes (symbiont-bearing vesicles or symbiosomes [98, 99]), but whether phagocytosis was the means of mitochondrial inclusion is not known yet. According to some hypotheses, the early host for mitochondria trapped its surface–contact partners in membrane protrusions [100, 101]. In case of a heterotrophic host capable of secreting extracellular digestive enzymes, such entrapment could serve as a poor-man’s phagocytosis [102]. A mixed vertical and horizontal transmission seems to be in effect in Burkholderia-infected Dictyostelium [103], indicative of facultative endosymbiosis.

Central control and organellogenesis

As partners become more dependent on each other, and as one party starts to dominate the other, central control evolves. Its ultimate form is the nuclear transfer of symbiont genes, requiring the presence of a nucleus and a mechanism to import proteins from the host cytosol to the symbiont. Evolved dependence on protein and lipid import mechanisms is a sign of endosymbiosis becoming irreversible.

For mitochondrial genes to undergo nuclear transfer, the host must have already been a (proto-)eukaryote. The ancestor of mitochondria could have been acquired before the nucleus, but only with the evolution of the true karyon could compartmentalized, safe transcription (safe from hybridization) be implemented. Symbiont genes relocating into the host nucleus are minimizing the effect of lower level of selection of the multilevel selection situation. With this step, eukaryotes left the prokaryotic domain for good.

After nuclear transfer of genes, it is necessary that proteins not produced by the symbiont anymore find their way back into the symbiont. Usually, this is a translocon-mediated protein import system installed by the host. With a protein import system in effect and a sufficient number of genes transferred to the nucleus, the symbiont could relinquish its protein-coding genes and protein-producing machinery, leveraging its genome. Moreover, this allows the host to introduce proteins of its own interest into the symbiont’s membrane.

The adenine nucleotide translocase (ANT) was probably introduced by the host into the mitochondrial membrane to exchange host-cytosolic ADP with symbiont ATP [36, 104]. ANT was most likely evolved within eukaryotes after the engulfment of the ancestral symbiont [105,106,107]. It was certainly in the host’s best interest to exploit the symbiont. If, however, it was the symbiont who invented ANT to give up ATP for the host, then any cheater bacterium capable of turning off its ANT while inside the host would have been under positive selection leading to the overpopulation of defecting symbionts, as was pointed out [108]. Group selection could have stabilized against cheaters, but only if a population of endosymbionts payed enough ATP to the host, so that host replicated faster (compared to other host cells) since the symbiont replicated with the host, the benefit was shared [109]. Other partner control mechanisms screening out cheaters are unknown yet.

No prokaryotic analogies of karyogenesis were found yet, though nuclear transfer is known in many eukaryotes [110]. Further features thought to be exclusive to mitochondrial and plastid integration have been recognized in more recent endosymbioses [71, 111, 112], drawing a picture of a continuum from symbiosis to organellogenesis (see Fig. 4). While nuclear integration renders the partnership obligate and irreversible, preventing the escape of the reduced partner, such mechanisms by no means represent an end state. They do not even ensure the survival of the symbiont, as amitochondriate eukaryotes attest. In the next section, we explore mechanisms that work against and could even ruin endosymbioses.


Engineering of plastid genomes

The invention of the particle gun provided a universal method for DNA delivery into living cells and subcellular compartments, including organelles. Stable transformation of the chloroplast genome by particle gun-mediated (biolistic) DNA delivery was first accomplished in the unicellular green alga C. reinhardtii (Boynton et al., 1988 ), followed by success in the seed plant species N. tabacum (tobacco Svab et al., 1990 Svab and Maliga, 1993 ). For more than 20 years, these two species have remained the organisms of choice for plastid transformation, and its application in both basic research and biotechnology. Although a few important crop species can now also be transformed (Sidorov et al., 1999 Ruf et al., 2001 Dufourmantel et al., 2004 ), progress with developing plastid transformation protocols for additional species has been somewhat slow, and many genetic model species and agriculturally relevant crops are still not transformable, notably including A. thaliana and all monocotyledonous species (Maliga, 2004 Maliga and Bock, 2011 Bock, 2014 ).

The integration of foreign DNA into the plastid genome occurs exclusively via homologous recombination, thus allowing very precise manipulations of the plastid genome, such as the introduction of point mutations at defined positions (Przibilla et al., 1991 Bock et al., 1994 ). The extraordinarily high activity of the homologous recombination system of plastids also facilitates the simultaneous modification of two distinct regions of the plastid genome by co-transformation experiments, in which two or more plasmid vectors are loaded on the microprojectiles for bombardment (Kindle et al., 1991 Carrer and Maliga, 1995 Krech et al., 2013 ). Amazingly, this approach can even be used to co-transform the nuclear genome (by non-homologous end joining) and the plastid genome (by homologous recombination) in a single experiment (Elghabi et al., 2011 ).

Over the years, plastid transformation in the two model systems, Chlamydomonas and tobacco, has become more and more routine, with the efficiency of plastid transformation now approaching that of nuclear transformation. This cannot be ascribed to a single methodological breakthrough, but rather is the result of many incremental improvements in the procedures involved in generating transplastomic cells and plants: the biolistic protocol, the transformation vectors, selectable markers and expression cassettes, and the tissue culture, selection and regeneration protocols. Along the way, a large toolkit for plastid genome engineering has been put together by both the tobacco and the Chlamydomonas communities (Maliga, 2004 Day and Goldschmidt-Clermont, 2011 Maliga and Bock, 2011 Bock, 2013 , 2014 ). This toolkit contains, for example, various selectable marker genes, reporter genes, and promoters and untranslated regions (5′- and 3′- UTRs) that confer widely different transgene expression levels. Recently, significant progress has also been made with improving plastid transgene expression in non-green plastid types, such as amyloplasts and chromoplasts (Valkov et al., 2011 Zhang et al., 2012 Caroca et al., 2013 ), and with developing methods for the inducible expression of plastid transgenes (Mühlbauer and Koop, 2005 Surzycki et al., 2007 Verhounig et al., 2010 ).

A salient feature of particle gun-mediated transformation is that DNA delivery is entirely based on a physical process. Thus, the biolistic method has no theoretical size limitation and large pieces of foreign DNA can be bombarded into the target compartment (Altpeter et al., 2005 ). So far, DNA pieces of up to 50 kb have been incorporated into the tobacco plastid genome (Adachi et al., 2007 ), and there is no reason to believe that much bigger pieces could not be introduced as well. Together with the small genome size (Figure 1) and the ease with which many genetic manipulations can be conducted, the capacity to accommodate large quantities of foreign DNA make the chloroplast an attractive target of synthetic biology. Based on pioneering work in microbial systems (Roodbeen and van Hest, 2009 Delaye and Moya, 2010 Cambray et al., 2011 ), two main branches of synthetic biology have emerged. Top-down synthetic biology approaches start from an existing biological system, and aim at reducing its complexity, ideally to a minimum-size system that consists of the smallest possible number of parts. Bottom-up synthetic biology approaches start with individual parts (building blocks) and try to construct artificial biological systems from first principles. The overarching goals of both approaches are very similar: (i) to further our understanding of the genetic elements and regulatory principles underlying functional biological systems and (ii) to design optimized biological systems for engineering applications. The latter goal brings synthetic biology in close proximity to biotechnology, and in fact many applications that nowadays come under the label of synthetic biology also could be viewed as advanced genetic engineering for biotechnological purposes (Peralta-Yahya et al., 2012 Paddon et al., 2013 ). This semantic issue notwithstanding, the amenability of plastids to large-scale genome manipulations with high precision facilitates both top-down and bottom-up approaches on the road to plant synthetic biology. In the following, the potential of plastids for synthetic biology is illustrated with two examples: (i) the design of minimum-size synthetic plastid genomes, a top-down approach and (ii) the build-up of new metabolic pathways in plastids via multigene engineering, a bottom-up approach.


CONCLUSION

Eukaryotes arose by the engulfment of prokaryotes and are thus genetic mosaics with two (animals and fungi) or three (plants) DNA-containing organelles. The integration of the third genetic compartment, the plastids, has led to photoautotrophic eukaryotes that are the nutritional basis for most life on earth. Plants had to evolve alternative means of metabolic coupling and organellar interaction hubs. We present a data set promoting the moss P. patens as a model organism for organelle biology, based on its evolutionary position and amenability to proteomics as well as microscopic studies. Comparative quantitative proteomics integrating validation on the single-protein level and metabolic pathway databases provides evidence that protein compartmentation and metabolic partitioning are highly flexible but well regulated in different kingdoms of life, different lineages within a kingdom, different tissues of a given species, between individual organelles of a single cell, and even at the suborganellar level by the formation of dynamic microcompartments in plastids and mitochondria.


Gray MW: Origin and evolution of organelle genomes. Curr Opin Genet Dev. 1993, 3: 884-890. 10.1016/0959-437X(93)90009-E.

Lake JA, Rivera MC: Was the nucleus the first endosymbiont?. Proc Natl Acad Sci USA. 1994, 91: 2880-2881. 10.1073/pnas.91.8.2880.

Martin W, Muller M: The hydrogen hypothesis for the first eukaryote. Nature. 1998, 392: 37-41. 10.1038/32096.

Margulis L, Dolan MF, Guerrero R: The chimeric eukaryote: origin of the nucleus from the karyomastigont in amitochondriate protists. Proc Natl Acad Sci USA. 2000, 97: 6954-6959. 10.1073/pnas.97.13.6954.

Gupta RS, Golding GB: The origin of the eukaryotic cell. Trends Biochem Sci. 1996, 21: 166-171. 10.1016/0968-0004(96)20013-1.

Martin W, Koonin EV: Introns and the origin of nucleus-cytosol compartmentalization. Nature. 2006, 440: 41-45. 10.1038/nature04531.

Zillig W, Palm P, Klenk H-P: A model of the early evolution of organisms: the arisal of the three domains of life from the common ancestor. The Origin and Evolution of the Cell. Edited by: Hartman H, Matsuno K. 1992, Singapore: World Scientific Publishing, 163-182.

McFadden GI: Mergers and acquisitions: malaria and the great chloroplast heist. Genome Biol. 2000, 1: reviews1026.1-1026.4. 10.1186/gb-2000-1-4-reviews1026.

Bhattacharya D, Yoon HS, Hackett JD: Photosynthetic eukaryotes unite: endosymbiosis connects the dots. Bioessays. 2004, 26: 50-60. 10.1002/bies.10376.

Keeling PJ: Diversity and evolutionary history of plastids and their hosts. Am J Botany. 2004, 91: 1481-1493.

Palmer JD: The symbiotic birth and spread of plastids: how many times and whodunit?. J Phycol. 2003, 39: 4-11. 10.1046/j.1529-8817.2003.02185.x.

Delwiche CF: Tracing the thread of plastid diversity through the tapestry of life. Am Nat. 1999, 154: S164-S177. 10.1086/303291.

Brinkman FS, Blanchard JL, Cherkasov A, Av-Gay Y, Brunham RC, Fernandez RC, Finlay BB, Otto SP, Ouellette BF, Keeling PJ, et al: Evidence that plant-like genes in Chlamydia species reflect an ancestral relationship between Chlamydiaceae, cyanobacteria, and the chloroplast. Genome Res. 2002, 12: 1159-1167. 10.1101/gr.341802.

Ludwig W, Klenk H-P: Overview: a phylogenetic backbone and taxonomic framework for prokaryotic systematics. Bergey's Manual of Systematic Bacteriology. Edited by: Boone DR, Garrity GM. 2001, Heidelberg, Germany: Springer, 1: 49-65.

Woese CR: Bacterial evolution. Microbiol Rev. 1987, 51: 221-271.

Everett KD: Chlamydia and Chlamydiales: more than meets the eye. Vet Microbiol. 2000, 75: 109-126. 10.1016/S0378-1135(00)00213-3.

Everett KD, Thao M, Horn M, Dyszynski GE, Baumann P: Novel chlamydiae in whiteflies and scale insects: endosymbionts 'Candidatus Fritschea bemisiae ' strain Falk and 'Candidatus Fritschea eriococci' strain Elm. Int J Syst Evol Microbiol. 2005, 55: 1581-1587. 10.1099/ijs.0.63454-0.

Horn M, Wagner M: Bacterial endosymbionts of free-living amoebae. J Eukaryot Microbiol. 2004, 51: 509-514. 10.1111/j.1550-7408.2004.tb00278.x.

Horn M, Wagner M: Evidence for additional genus-level diversity of Chlamydiales in the environment. FEMS Microbiol Lett. 2001, 204: 71-74. 10.1111/j.1574-6968.2001.tb10865.x.

Horn M, Collingro A, Schmitz-Esser S, Beier CL, Purkhold U, Fartmann B, Brandt P, Nyakatura GJ, Droege M, Frishman D, et al: Illuminating the evolutionary history of chlamydiae. Science. 2004, 304: 728-730. 10.1126/science.1096330.

Stephens RS, Kalman S, Lammel C, Fan J, Marathe R, Aravind L, Mitchell W, Olinger L, Tatusov RL, Zhao Q, et al: Genome sequence of an obligate intracellular pathogen of humans: Chlamydia trachomatis. Science. 1998, 282: 754-759. 10.1126/science.282.5389.754.

Greub G, Raoult D: History of the ADP/ATP-translocase-encoding gene, a parasitism gene transferred from a Chlamydiales ancestor to plants 1 billion years ago. Appl Environ Microbiol. 2003, 69: 5530-5535. 10.1128/AEM.69.9.5530-5535.2003.

Royo J, Gimez E, Hueros G: CMP-KDO synthetase: a plant gene borrowed from gram-negative eubacteria. Trends Genet. 2000, 16: 432-433. 10.1016/S0168-9525(00)02102-8.

Ortutay C, Gaspari Z, Toth G, Jager E, Vida G, Orosz L, Vellai T: Speciation in Chlamydia: genomewide phylogenetic analyses identified a reliable set of acquired genes. J Mol Evol. 2003, 57: 672-680. 10.1007/s00239-003-2517-3.

Linka N, Hurka H, Lang BF, Burger G, Winkler HH, Stamme C, Urbany C, Seil I, Kusch J, Neuhaus HE: Phylogenetic relationships of non-mitochondrial nucleotide transport proteins in bacteria and eukaryotes. Gene. 2003, 306: 27-35. 10.1016/S0378-1119(03)00429-3.

Amiri H, Karlberg O, Andersson SG: Deep origin of plastid/parasite ATP/ADP translocases. J Mol Evol. 2003, 56: 137-150. 10.1007/s00239-002-2387-0.

Schmitz-Esser S, Linka N, Collingro A, Beier CL, Neuhaus HE, Wagner M, Horn M: ATP/ADP translocases: a common feature of obligate intracellular amoebal symbionts related to Chlamydiae and Rickettsiae. J Bacteriol. 2004, 186: 683-691. 10.1128/JB.186.3.683-691.2004.

Wolf YI, Aravind L, Koonin EV: Rickettsiae and Chlamydiae: evidence of horizontal gene transfer and gene exchange. Trends Genet. 1999, 15: 173-175. 10.1016/S0168-9525(99)01704-7.

Rodriguez-Ezpeleta N, Brinkmann H, Burey SC, Roure B, Burger G, Loffelhardt W, Bohnert HJ, Philippe H, Lang BF: Monophyly of primary photosynthetic eukaryotes: green plants, red algae, and glaucophytes. Curr Biol. 2005, 15: 1325-1330. 10.1016/j.cub.2005.06.040.

Huang J, Gogarten JP: Ancient horizontal gene transfer can benefit phylogenetic reconstruction. Trends Genet. 2006, 22: 361-366. 10.1016/j.tig.2006.05.004.

deBary HA: The Phenomenon of Symbiosis [in German]. 1879, Strassburg, Germany: Verlag von Karl J. Trubner

Matsuzaki M, Misumi O, Shin IT, Maruyama S, Takahara M, Miyagishima SY, Mori T, Nishida K, Yagisawa F, Nishida K, et al: Genome sequence of the ultrasmall unicellular red alga Cyanidioschyzon merolae 10D. Nature. 2004, 428: 653-657. 10.1038/nature02398.

O'Brien EA, Koski LB, Zhang Y, Yang L, Wang E, Gray MW, Burger G, Lang BF: TBestDB: a taxonomically broad database of expressed sequence tags (ESTs). Nucleic Acids Res. 2007, D445-D451. 10.1093/nar/gkl770. 35 Database

Emanuelsson O, Nielsen H, von Heijne G: ChloroP, a neural network-based method for predicting chloroplast transit peptides and their cleavage sites. Protein Sci. 1999, 8: 978-984.

Emanuelsson O, Nielsen H, Brunak S, von Heijne G: Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol. 2000, 300: 1005-1016. 10.1006/jmbi.2000.3903.

Collingro A, Toenshoff ER, Taylor MW, Fritsche TR, Wagner M, Horn M: 'Candidatus Protochlamydia amoebophila ', an endosymbiont of Acanthamoeba spp. Int J Syst Evol Microbiol. 2005, 55: 1863-1866. 10.1099/ijs.0.63572-0.

Weisburg WG, Hatch TP, Woese CR: Eubacterial origin of chlamydiae. J Bacteriol. 1986, 167: 570-574.

Nelson KE, Paulsen IT, Heidelberg JF, Fraser CM: Status of genome projects for nonpathogenic bacteria and archaea. Nat Biotechnol. 2000, 18: 1049-1054. 10.1038/80235.

Brochier C, Philippe H, Moreira D: The evolutionary history of ribosomal protein RpS14: horizontal gene transfer at the heart of the ribosome. Trends Genet. 2000, 16: 529-533. 10.1016/S0168-9525(00)02142-9.

Gogarten JP, Townsend JP: Horizontal gene transfer, genome innovation and evolution. Nat Rev Microbiol. 2005, 3: 679-687. 10.1038/nrmicro1204.

Martin W, Herrmann RG: Gene transfer from organelles to the nucleus: how much, what happens, and why?. Plant Physiol. 1998, 118: 9-17. 10.1104/pp.118.1.9.

Gogarten JP, Doolittle WF, Lawrence JG: Prokaryotic evolution in light of gene transfer. Mol Biol Evol. 2002, 19: 2226-2238.

Ochman H, Lawrence JG, Groisman EA: Lateral gene transfer and the nature of bacterial innovation. Nature. 2000, 405: 299-304. 10.1038/35012500.

Jain R, Rivera MC, Lake JA: Horizontal gene transfer among genomes: the complexity hypothesis. Proc Natl Acad Sci USA. 1999, 96: 3801-3806. 10.1073/pnas.96.7.3801.

Keeling PJ, Burger G, Durnford DG, Lang BF, Lee RW, Pearlman RE, Roger AJ, Gray MW: The tree of eukaryotes. Trends Ecol Evol. 2005, 20: 670-676. 10.1016/j.tree.2005.09.005.

Baldauf SL, Roger AJ, Wenk-Siefert I, Doolittle WF: A kingdom-level phylogeny of eukaryotes based on combined protein data. Science. 2000, 290: 972-977. 10.1126/science.290.5493.972.

Douzery EJ, Snell EA, Bapteste E, Delsuc F, Philippe H: The timing of eukaryotic evolution: does a relaxed molecular clock reconcile proteins and fossils?. Proc Natl Acad Sci USA. 2004, 101: 15386-15391. 10.1073/pnas.0403984101.

Hedges SB, Blair JE, Venturi ML, Shoe JL: A molecular timescale of eukaryote evolution and the rise of complex multicellular life. BMC Evol Biol. 2004, 4: 2-10.1186/1471-2148-4-2.

Huang J, Mullapudi N, Lancto CA, Scott M, Abrahamsen MS, Kissinger JC: Phylogenomic evidence supports past endosymbiosis, intracellular and horizontal gene transfer in Cryptosporidium parvum. Genome Biol. 2004, 5: R88-10.1186/gb-2004-5-11-r88.

Berriman M, Ghedin E, Hertz-Fowler C, Blandin G, Renauld H, Bartholomeu DC, Lennard NJ, Caler E, Hamlin NE, Haas B, et al: The genome of the African trypanosome Trypanosoma brucei. Science. 2005, 309: 416-422. 10.1126/science.1112642.

Archibald JM, Rogers MB, Toop M, Ishida K, Keeling PJ: Lateral gene transfer and the evolution of plastid-targeted proteins in the secondary plastid-containing alga Bigelowiella natans. Proc Natl Acad Sci USA. 2003, 100: 7678-7683. 10.1073/pnas.1230951100.

Esser C, Ahmadinejad N, Wiegand C, Rotte C, Sebastiani F, Gelius-Dietrich G, Henze K, Kretschmann E, Richly E, Leister D, et al: A genome phylogeny for mitochondria among alpha-proteobacteria and a predominantly eubacterial ancestry of yeast nuclear genes. Mol Biol Evol. 2004, 21: 1643-1660. 10.1093/molbev/msh160.

Martin W, Rujan T, Richly E, Hansen A, Cornelsen S, Lins T, Leister D, Stoebe B, Hasegawa M, Penny D: Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc Natl Acad Sci USA. 2002, 99: 12246-12251. 10.1073/pnas.182432999.

Reyes-Prieto A, Hackett JD, Soares MB, Bonaldo MF, Bhattacharya D: Cyanobacterial contribution to algal nuclear genomes is primarily limited to plastid functions. Curr Biol. 2006, 16: 2320-2325. 10.1016/j.cub.2006.09.063.

Gardner MJ, Hall N, Fung E, White O, Berriman M, Hyman RW, Carlton JM, Pain A, Nelson KE, Bowman S, et al: Genome sequence of the human malaria parasite Plasmodium falciparum. Nature. 2002, 419: 498-511. 10.1038/nature01097.

Loftus B, Anderson I, Davies R, Alsmark UC, Samuelson J, Amedeo P, Roncaglia P, Berriman M, Hirt RP, Mann BJ, et al: The genome of the protist parasite Entamoeba histolytica. Nature. 2005, 433: 865-868. 10.1038/nature03291.

Molmeret M, Horn M, Wagner M, Santic M, Abu Kwaik Y: Amoebae as training grounds for intracellular bacterial pathogens. Appl Environ Microbiol. 2005, 71: 20-28. 10.1128/AEM.71.1.20-28.2005.

Winkler HH, Neuhaus HE: Non-mitochondrial ATP transport. Trends Biochem Sci. 1999, 24: 64-68. 10.1016/S0968-0004(98)01334-6.

Tjaden J, Winkler HH, Schwoppe C, Van Der Laan M, Mohlmann T, Neuhaus HE: Two nucleotide transport proteins in Chlamydia trachomatis, one for net nucleoside triphosphate uptake and the other for transport of energy. J Bacteriol. 1999, 181: 1196-1202.

Toyota K, Tamura M, Ohdan T, Nakamura Y: Expression profiling of starch metabolism-related plastidic translocator genes in rice. Planta. 2006, 223: 248-257. 10.1007/s00425-005-0128-5.

Archibald JM: Jumping genes and shrinking genomes: probing the evolution of eukaryotic photosynthesis with genomics. IUBMB Life. 2005, 57: 539-547.

Fast NM, Kissinger JC, Roos DS, Keeling PJ: Nuclear-encoded, plastid-targeted genes suggest a single common origin for apicomplexan and dinoflagellate plastids. Mol Biol Evol. 2001, 18: 418-426.

Ryall K, Harper JT, Keeling PJ: Plastid-derived type II fatty acid biosynthetic enzymes in chromists. Gene. 2003, 313: 139-148. 10.1016/S0378-1119(03)00671-1.

Nakabachi A, Yamashita A, Toh H, Ishikawa H, Dunbar HE, Moran NA, Hattori M: The 160-kilobase genome of the bacterial endosymbiont Carsonella. Science. 2006, 314: 267-10.1126/science.1134196.

Heldt H-W: Plant Biochemistry. 2005, Burlington, MA: Elsevier Inc, 3

Lange BM, Croteau R: Isopentenyl diphosphate biosynthesis via a mevalonate-independent pathway: isopentenyl monophosphate kinase catalyzes the terminal enzymatic step. Proc Natl Acad Sci USA. 1999, 96: 13714-13719. 10.1073/pnas.96.24.13714.

Page JE, Hause G, Raschke M, Gao W, Schmidt J, Zenk MH, Kutchan TM: Functional analysis of the final steps of the 1-deoxy-D-xylulose 5-phosphate (DXP) pathway to isoprenoids in plants using virus-induced gene silencing. Plant Physiol. 2004, 134: 1401-1413. 10.1104/pp.103.038133.

Lange BM, Rujan T, Martin W, Croteau R: Isoprenoid biosynthesis: the evolution of two ancient and distinct pathways across genomes. Proc Natl Acad Sci USA. 2000, 97: 13172-13177. 10.1073/pnas.240454797.

Marin B, Nowack EC, Melkonian M: A plastid in the making: evidence for a second primary endosymbiosis. Protist. 2005, 156: 425-432. 10.1016/j.protis.2005.09.001.

Weber AP, Linka M, Bhattacharya D: Single, ancient origin of a plastid metabolite translocator family in Plantae from an endomembrane-derived ancestor. Eukaryot Cell. 2006, 5: 609-612. 10.1128/EC.5.3.609-612.2006.

Stiller JW, Hall BD: The origin of red algae: implications for plastid evolution. Proc Natl Acad Sci USA. 1997, 94: 4520-4525. 10.1073/pnas.94.9.4520.

Stiller JW, Riley J, Hall BD: Are red algae plants? A critical evaluation of three key molecular data sets. J Mol Evol. 2001, 52: 527-539.

Frickey T, Lupas AN: PhyloGenie: automated phylome generation and analysis. Nucleic Acids Res. 2004, 32: 5231-5238. 10.1093/nar/gkh867.

Edgar RC: MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 2004, 32: 1792-1797. 10.1093/nar/gkh340.

Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG: The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997, 25: 4876-4882. 10.1093/nar/25.24.4876.

Guindon S, Gascuel O: A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol. 2003, 52: 696-704. 10.1080/10635150390235520.

Felsenstein J: PHYLIP (Phylogeny Inference Package) version 3.65 (distributed by the author). 2005, Seattle, Washington: Department of Genome Sciences, University of Washington

Schmidt HA, Strimmer K, Vingron M, von Haeseler A: TREE-PUZZLE: maximum likelihood phylogenetic analysis using quartets and parallel computing. Bioinformatics. 2002, 18: 502-504. 10.1093/bioinformatics/18.3.502.


References

Palmer JD. Comparative organization of chloroplast genomes. Annu Rev Genet. 198519:325–54.

Wicke S, Schneeweiss GM, de Pamphilis CW, Müller KF, Quandt D. The evolution of the plastid chromosome in land plants: gene content, gene order, gene function. Plant Mol Biol. 201176:273—97.

Palmer JD. Molecular evolution: a single birth of all plastids? Nature. 2000405:32–3.

McFadden GI, van Dooren GG. Evolution: red algal genome affirms a common origin of all plastids. Curr Biol. 2004R514:516.

Keeling PJ, et al. Philos Trans R Soc Lond Ser B Biol Sci. 2010365:729–48.

Martin W, Rujan T, Richly E, Hansen A, Cornelsen S, Lins T, Leister D, Stoebe B, Hasegawa M, Penny D. Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc Natl Acad Sci U S A. 200299:12246–51.

Raubeson LA, Jansen RK. Chloroplast genomes of plants. In: Henry RJ, editor. Plant diversity and evolution: genotypic and phenotypic variation in higher plants. Wallingford, UK: CABI Publishing 2005. p. 45–68.

Kolodner R, Tewari KK. Inverted repeats in chloroplast DNA from higher plants. P Natl Acad Sci USA. 197976:41–5.

Maréchal A, Parent J, Véronneau-Lafortune F, Joyeux A, Lang BF, Brisson N. Whirly proteins maintain plastid genome stability in Arabidopsis. Proc Natl Acad Sci U S A. 2009106:14693–8.

Palmer JD. Plastid chromosomes: structure and evolution. In: Bogorad L, Vasil IK, editors. Cell culture and somatic genetics of plant, vol 7A. Molecular biology of plastids. San Diego: Academic Press 1991. p. 5—53.

Sugiura M. The chloroplast genome. Plant Mol Biol. 199219:149–68.

Bock R. Structure, function, and inheritance of plastid genomes. In: Bock R, editor. Cell and molecular biology of plastids. Berlin Heidelberg, 63: Springer. p. 29.

dePamphilis CW, Palmer JD. Loss of photosynthetic and chlororespiratory genes from the plastid genome of a parasitic flowering plant. Nature. 1990348:337–9.

Wolfe KH, Morden CW, Palmer JD. Function and evolution of a minimal plastid genome from a nonphotosynthetic parasitic plant. Proc Natl Acad Sci U S A. 199289:10648–52.

Nickrent DL, Ouyang R, Joel D, dePamphilis CW. Do nonasterid holoparasitic flowering plants have plastid genomes? Plant Mol Biol. 199734:717–29.

Funk H, Berg S, Krupinska K, Maier U, Krause K. Complete DNA sequences of plastid genomes of two parasitic flowering plant species Cuscuta reflexa and Cuscuta gronovii. BMC Plant Biol. 20077:45.

McNeal JR, Kuehl J, Boore J, dePamphilis C. Complete plastid genome sequences suggest strong selection for retention of photosynthetic genes in the parasitic plant genus Cuscuta. BMC Plant Biol. 20077:57.

Krause K. From chloroplasts to “cryptic” plasids: evolution of plastid genomes in parasitic plants. Curr Genet. 200854:111–21.

Wicke S, Müller KF, dePamphilis CW, Quandt D, Wickett NJ, Zhang Y, Renner SS, Schneeweiss GM. Mechanisms of functional and physical genome reduction in photosynthetic and nonphotosynthetic parasitic plants of the broomrape family. Plant Cell. 201325:3711–25.

Wicke S, Müller KF, de Pamphilis CW, Quandt D, Bellot S, Schneeweiss GM. Mechanistic model of evolutionary rate variation en route to a nonphotosynthetic lifestyle in plants. P Natl A Sci. 2016113:9045–50.

Samigullin TH, Logacheva MD, Penin AA, Vallejo CM. Complete plastid genome of the recent holoparasites Lathraea squamaria reveals earliest stages of plastome reduction in Orobanchaceae. PLoS One. 2016 https://doi.org/10.1371/journal.pone.0150718.

Delavault PM, Russo NM, Lusson NA, Thalouarn P. Organization of the reduced plastid genome of Lathraea clandestina an achlorophyllous parasitic plant. Physiol Plant. 199696:674–82.

Wickett NJ, Zhang Y, Hansen SK, Roper JM, Kuehl JV, Plock SA, Wolf PG, dePamphilis CW, Boore JL, Goffinet B. Functional gene losses occur with minimal size reduction in the plastid genome of the parasitic liverwort Aneura mirabilis. Mol Biol Evol. 200825:393–401.

Molina J, Hazzouri KM, Nickrent D, Geisler M, Meyer RS, Pentony MM, Flowers JM, Pelser P, Barcelona J, Inovejas SA, et al. Possible loss of the chloroplast genome in the parasitic flowering plant Rafflesia lagascae (Rafflesiaceae). Mol Biol Evol. 201431:793–803.

Naumann J, Der JP, Wafula EK, Jones SS, Wagner ST, Honaas LA, Ralph PE, Bolin JF, Maass E, Neinhuis C, et al. Detecting and characterizing the highly divergent plastid genome of the nonphotosynthetic parasitic plant Hydnora visseri (Hydnoraceae). Genome Biol Evol. 20168:345–63.

Bellot S, Renner SS. The plastomes of two species in the endoparasite genus Pilostyles (Apodanthaceae) each retain just five or six possibly functional genes. Genome Biol Evol. 20158:189–201.

Roquet C, Coissac É, Cruaud C, Boleda M, Boyer F, Alberti A, Gielly L, Taberlet P, Thuiller W, Van Es J, Lavergne S. Understanding the evolution of holoparasitic plants: the complete plastid genome of the holoparasites Cytinus hypocistis (Cytinaceae). Ann Bot. 2016118:885–96.

Zhang R, Wang J, Han K, Ren T, Zeng S, Biffin E, Liu Z. Complete chloroplast genome sequence of Pedicularis cheilanthifolia, an alpine plant in China. Conserv Genet Resour. 2017doi:https://doi.org/10.1007/s12686-017-0740-2

Downie SR, Palmer JD. Restriction site mapping of the chloroplast DNA inverted repeat: a molecular phylogeny of the Asteridae. Ann Mo Bot Gard. 199279:266–83.

Perry AS, Wolfe KH. Nucleotide substitution rates in legume chloroplast DNA depend on the presence of the inverted repeat. J Mol Evol. 200255:501–8.

Olmstead RG, dePamphilis CW, Wolfe AD, Young ND, Elisons WJ, Reeves PA. Disintegration of the Scrophulariaceae. Am J Bot. 200188:348–61.

McNeal JR, Bennet JR, Wolfe AD, Mathews S. Phylogeny and origins of holoparasitism in Orobanchaceae. Am J Bot. 2013100:971–83.

Estep MC, Gowda BS, Huang K, Timko MP, Bennetzen JL. Genomic characterization for parasitic weeds of the genus by sample sequence analysis. Plant Genome. 20125:30–41.

Doyle JJ, Doyle JL. A rapid DNA isolation procedure for small quantities of fresh leaf tissue. Phytochem Bull. 198719:11–5.

Vaughn JN, Chaluvadi SR, Tushar RL, Bennetzen JL. Whole plastome sequences from five ginger species facilitate marker development and define limits to barcode methodology. PLoS One. 2014 https://doi.org/10.1371/journal.pone.0108581.

Darling ACE, Mau B, Blattner FR, Perna NT. Mauve: multiple alignment of conserved genomic sequence with rearrangements. Genet Res. 200414:394–1403.

Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S, Buxton S, Cooper A, Markowitz S, Duran C, Thierer T, Ashton B, Mentjies P, Drummond A. Geneious basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics. 201228:1647–9.

Cui L, Veeraraghavan N, Richter A, Wall K, Jansen RK, Leebens-Mack J, Makalowska I, dePamphilis CW. ChloroplastDB: the chloroplast genome database. Nucleic Acids Res. 200634:D692–6.

Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG. Clustal W and Clustal X version 2.0. Bioinformatics. 200723:2947–8.

Kumar S, Stecher G, Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0. Mol Biol Evol. 201633:1870–4.

Ronquist F, Huelsenback JP. MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 200319:1572–4.

Westwood JH, Yoder JI, Timko MP, dePamphilis CW. The evolution of parasitism in plants. Trends Plant Sci. 201015:227–35.

Bremer K, Fijs EM, Bremer B. Molecular phylogenetic dating of asteroid flowering plants shows early cretaceous diversification. Syst Biol. 200453:496–505.

Wolfe AD, Randle CP, Liu L, Steiner KE. Phylogeny and biogeography of Orobanchaceae. Folia Geobot. 200540:115–34.

Naumann J, Salomo K, Der JP, Wafula EK, Bolin JF, Maass E, Frenzke L, Samain MS, Neinhuis C, dePamphilis CW, Wanke S. Single-copy nuclear genes place haustorial Hydnoraceae within Piperales and reveal a cretaceous origin of multiple parasitic angiosperm lineages. PLoS One. 2013 https://doi.org/10.1371/journal.pone.0079204.

Wolfe KH, Li WH, Sharp PM. Rates of nucleotide substitution vary greatly among plant mitochondrial, chloroplast, and nuclear DNAs. Proc Natl Acad Sci U S A. 198784:9054–8.

Goulding SE, Wolfe KH, Olmstead RG, Morden CW. Ebb and flow of the chloroplast inverted repeat. Mol Gen Genet. 1996252:195–206.

Wang RJ, Cheng CL, Chang CC, Wu CL, Su TM, Chaw SM. Dynamics and evolution of the inverted repeat-large single copy junctions in the chloroplast genomes of monocots. BMC Evol Biol. 20088:36.

Downie SR, Jansen RK. A comparative analysis of whole plastid genomes from the Apiales: expansion and contraction of the inverted repeat, mitochondrial to plastid transfer of DNA, and identification of highly divergent noncoding regions. Syst Bot. 201540:336–51.

Knox EB, Palmer JD. The chloroplast genome arrangement of Lobelia thuliniana (Lobeliaceae): expansion of the inverted repeat in an ancestor of the Campanulales. Plant Syst Evol. 1999214:49–64.

Dugas DV, Hernandez D, Koenen EJM, Schwarz E, Straub S, Hughes CE, Jansen RK, Nageswara-Rao M, Staats M, Trujillo JT, Hajrah NH, Alharbi NS, Al-Malki AL, Sabir JSM, Bailey CD. Mimosoid legume plastome evolution: IR expansion, tandem repeat expansions, and accelerated rate of evolution in clpP. Sci Rep. 20155:16958.

Zhu A, Guo W, Gupta S, Fan W, Mower JP. Evolutionary dynamics of the plastid inverted repeat: the effects of expansion, contraction, and loss on substitution rates. New Phytol. 2016209:1747–56.

Chumley TW, Palmer JD, Mower JP, Fourcade HM, Calie PJ, Boore JL, Jansen RK. The complete chloroplast genome sequence of Pelargonium x hortorum: organization and evolution of the largest and most highly rearranged chloroplast genome of land plants. Mol Biol Evol. 200623:2175–90.

Casano LM, Zapata JM, Marti M, Sabater B. Chlororespiration and poising of cyclic electron transport. J Biol Chem. 2000275:942–8.

Nixon PJ. Chlororespiration. Philos Trans R Soc B Biol Sci. 2000355:1541–7.

Martín M, Funk HT, Serrot PH, Poltnigg P, Sabater B. Functional characterization of the thylakoid Ndh complex phosphorylation by site-directed mutations in the ndhF gene. BBA-Bioenergetics. 20091787:920–8.

Martín M, Sabater B. Plastid ndh genes in plant evolution. Plant Physiol Biochem. 201048:636–45.

Friedrich T, Steinmüller K, Weiss H. The proton-pumping respiratory complex I of bacteria and mitochondria and its homologue in chloroplasts. FEBS Lett. 1995367:107–11.

McCoy SR, Kuehl JV, Boore JL, Raubeson LA. The complete plastid genome sequence of Welwitschia mirabilis: an unusually compact plastome with accelerated divergence rates. BMC Evol Biol. 20088:130.

Wu C, Lai Y, Lin C, Wang Y, Chaw S, et al. Mol Phylogenet Evol. 200952:115–24.

Chang CC, Lin HC, Lin IP, Chow TY, Chen HH, Chen WH, Cheng CH, Lin CY, Liu SM, Chang CC, Chaw SM. The chloroplast genome of Phalaenopsis aphrodite (Orchidaceae): comparative analysis of evolutionary rate with that of grasses and its phylogenetic implications. Mol Biol Evol. 200623:279–91.

Wu F, Chan M, Liao D, Hsu C, Lee Y, Daniell H, Duvall M, Lin C. Complete chloroplast genome of Oncidium Gower Ramsey and evaluation of molecular markers for identification and breeding in Oncidiinae. BMC Plant Biol. 201010:68.

Barrett CF, Freudenstein JV, Li J, Mayfield-Jones DR, Perez L, Pires JC, Santos C. Investigating the path of plastid genome degradation in an early-transitional clade of heterotrophic orchids and implications for heterotrophic angiosperms. Mol Biol Evol. 201431:3095–112.

de Vries J, Sousa FL, Bölter B, Soll J, Gould SB. YCF1: a green TIC? Plant Cell. 201527:1827–33.

Guisinger MM, Chumley TW, Kuehl JV, Boore JL, Jansen RK. Implications of the plastid genome sequence of typha (typhaceae, poales) for understanding genome evolution in poaceae. J Mol Evol. 201070:146–66.

Logacheva MD, Schelkunov MI, Shtratnikova VY, Matveeva MV, Penin AA. Comparative analysis of plastid genomes of non-photosynthetic Ericaceae and their photosynthetic relatives. Sci Report. 2016 https://doi.org/10.1038/srep30042.

Oliver M, Murdock A, Mishler BD, Kuehl J, Boore J, Mandoli D, Everett K, Wolf PG, Duffy A, Karol KG. Chloroplast genome sequence of the moss Tortula ruralis: gene content, polymorphism, and structural arrangement relative to other green plant chloroplast genomes. BMC Genomics. 201011:143.

Wolf PG, Der J, Duffy A, Davidson J, Grusz A, Pryer KM. The evolution of chloroplast genes and genomes in ferns. Plant Mol Biol. 2010 https://doi.org/10.1007/s11103-010-9706-4.

Downie SR, Katz-Downie DS, Wolfe KH, Calie PJ, Palmer JD. Structure and evolution of the largest chloroplast gene (ORF2280): internal plasticity and multiple gene loss during angiosperm evolution. Curr Genet. 199425:367–3781.

Jansen RK, Cai Z, Raubeson LA, Daniell H, de Pamphilis CW, Leebens-Mack JH, Müller KF, Guisinger-Bellian M, Haberle RC, Hansen AK, Chumley TW, Lee SB, Peery R, JR MN, Kuehl JV, Boore JL. Analysis of 81 genes from 64 plastid genomes resolves relationships in angiosperms and identifies genome-scale evolutionary patterns. Proc Natl Acad Sci U S A. 2007104:19369–74.

Nakkaew A, Chotigeat W, Eksomtramage T, Phongdara A. Cloning and expression of a plastid-encoded subunit betacarboxyltransferase gene (accD) and a nuclear-encoded subunit biotin carboxylase of acetyl-CoA carboxylase from oil palm (Elaeis guineensis Jacq.). Plant Sci. 2008175:497–504.

Bennet JR, Mathews S. Phylogeny of the parasitic plant family Orobanchaceae inferred from phytochrome A. Am J Bot. 200693:1039–51.

Palmer JD. Chloroplast DNA evolution and biosystematics uses of chloroplast DNA variation. Am Nat. 1987130:S6–S29.

Tsudzuki J, Nakashima K, Tsudzuki T, Hiratsuka J, Shibata M, Wakasugi T, Sugiura M. Chloroplast DNA of black pine retains a residual inverted repeat lacking rRNA genes: nucleotide sequences of trnQ, trnK, psbA, trnI and trnH and the absence of rps16. Mol Gen Genet. 1992232:206—14.

Plunkett GM, Downie SR. Expansion and contraction of the chloroplast inverted repeat in Abiaceae subfamily Apioideae. Syst Bot. 2000:648–67.

Daniell H, Lee S, Grevich J, Saski C, Quesada-Vargas T, Guda C, Tomkins J, Jansen RK. Complete chloroplast genome sequences of Solanum bulbocastanum, Solanum lycopersicum and comparative analyses with other Solanaceae genomes. Theor Appl Genet. 2006112:1503–18.

Wolfe PG, Roper JM, Duffy AM. The evolution of chloroplast genome structure in ferns. Genome. 201053:731–8.

Grewe F, Viehoever P, Weisshaar B, Knoop V. A trans-splicing group I intron and tRNA-hyperediting in the mitochondrial genome of the lycophyte Isoëtes engelmannii. Nucleic Acids Res. 200937:5093–104.

Guo W, Grewe F, Cobo-Clark A, Fan W, Duan Z, Adams RP, Schwarzbach AE, Mower JP. Predominant and substoichiometric isomers of the plastid genome coexist within Juniperus plants and have shifted multiple times during cupressophyte evolution. Genome Biol Evol. 20146:580–90.

Svab Z, Hajdukiewicz P, Maliga P. Stable transformation of plastids in higher plants. Proc Natl Acad Sci U S A. 199087:8526–30.

Sikdar SR, Serino G, Chaudhuri S, Maliga P. Plastid transformation in Arabidopsis thaliana. Plant Cell Rep. 199818:20–4.

Sidorov VA, Kasten D, Pang SZ, Hajdukiewicz PTJ, Staub JM, Nehra NS. Stable chloroplast transformation in potato: use of green fluorescent protein as a plastid marker. Plant J. 199919:209–16.

Wani SH, Haider N, Kumar H, Singh NB. Plant plastid engineering. Curr Genomics. 201011:500–12.


Background

While the primary acquisition of the plastid from a free-living cyanobacterium is believed to have occurred only once [1], plastids have continued to spread through eukaryotes by means of secondary and tertiary endosymbiosis. This is the process whereby a plastid-containing, free-living eukaryote is consumed by another eukaryotic cell and becomes an organelle itself. Primary plastids (exemplified by those of plants) have two membranes, while secondary plastids have additional membranes corresponding to the outer membrane of the engulfed eukaryote and the phageosomal membrane of the host, as well as the original membranes of the primary plastid [2, 3], although in some lineages membranes have subsequently been lost. The nucleus of the engulfed cell is, in all but two described cases, absent, and the genes encoding plastid-targeted proteins having been relocated to the host nucleus [4–6]. The exceptions are the cryptomonads and chlorarachniophytes, which contain nucleomorphs, the remnant nuclei of the plastid-containing algae that were engulfed in the secondary endosymbioses that gave rise to these lineages (Figure 1). The cryptomonad endosymbiont is derived from a red alga, while that of chlorarachniophytes is derived from a green alga. Their genomes encode very few genes, and most of them are housekeeping genes for replication, transcription and protein folding and degradation [7, 8]. A handful of proteins related to plastid function have also been retained, however, they are relatively few [7, 9, 10]. The periplastidial space (equivalent to the cytosol of the engulfed alga) itself has specific metabolic processes, such as starch synthesis in cryptomonads, but most of the proteins for these pathways are missing from the nucleomorph genome [7] and are anticipated to be found in the nuclear genome, as has been shown for a few examples [11].

Endosymbiotic events that gave rise to cryptomonads and chlorarachniophytes.

The nucleomorph is often thought of as an anomaly, a rare occurrence, since it is known only in cryptomonads and chlorarachniophytes, but if one considers 'loss or gain' rather than 'presence or absence' then it is perhaps not so anomalous. All lineages that are known to contain secondary plastids (haptophytes, heterokonts, cryptomonads, dinoflagellates, apicomplexans, euglenids and chlorarachniophytes) have ancestors that contained a nucleomorph. Depending on the number of secondary endosymbiotic events that took place, which is still contentious [3, 12–14], the number of nucleomorph losses and gains differs. The balance of molecular evidence points to two events involving green algae [15, 16] and one involving a red alga [17–19]. With respect to green algae this means one lineage lost its nucleomorph and one retained it. With respect to red algae, this means a single nucleomorph gain (if one accepts the chromalveolate hypothesis [20]) and perhaps only one loss, if cryptomonads are the deepest branch of chromalveolates, or perhaps two if they diverged later. Overall, lineages retaining nucleomorphs may be as common as lineages that lost them, or at least the proportions are very similar. Whatever the case, nucleomorphs existed in the common ancestors of a great deal of algal diversity, so the study of the lineages in which they remain may help us understand the process of secondary (and higher order) endosymbiotic events, especially the reduction and subsequent loss of the enslaved genome.

Cryptomonads and chlorarachniophytes arose from separate endosymbiotic events, and neither host cell nor endosymbiont are very closely related. Yet the nucleomorph genomes of the cryptomonad, Guillardia theta [7] and the chlorarachniophyte, Bigelowiella natans [8–10] share several characteristics. Both nucleomorph genomes have undergone substantial gene loss and are ultra-compact compared to their free-living relatives in the red and green algae. Some of these features, such as overlapping genes, short intergenic regions, a reduction in elements like transposons, and the presence of multigene transcripts have been found in other compact eukaryotic genomes such as microsporidia [21, 22]. Compact genomes and many of these features are common to endosymbionts in general, however, until the sequences of the G. theta and B. natans, nucleomorph genomes were completed, all known endosymbiont genomes have been of prokaryotic origin. The best examples of prokaryotic endosymbiont genomes are those of the mitochondrion, once a free-living alpha-proteobacterium, and the chloroplast, once a free-living cyanobacterium [1]. Also well described, although not organellar, are the bacterial endosymbionts of insects, of these there are several complete genomes for example, Wolbachia [23–25], Buchnera [26], Wigglesworthia [27] and Blochmania [28], the features of which have been compared and defined [29–31]. These bacteria reside within a range of diverse insects but, while they retain certain distinct genes that can be linked to the physiology of their host, they show similar patterns of genome reduction, strong mutational AT bias and strict amino acid bias at high expression genes [32] an effect of selection against mutation driven amino acid changes [31, 33]. The AT mutational pressure in endosymbionts, is sometimes very extreme estimated to be a remarkable 90% GC->AT in Buchnera [34]. A universal AT mutational bias, has been suggested because many types of spontaneous mutations (e.g. the deamination of cytosine) cause GC to AT changes [35]. The effects of this mutational bias may be more pronounced and gene loss more rapid in small, endosymbiont genomes because they are deficient in at least one DNA repair mechanism, experience strong genetic drift and have experienced a relaxation of selection in the intracellular environment in comparison to free-living existence [31, 33].

There is less chromosomal information for eukaryotic obligate intracellular parasites, however certain alveolate and microsporidian genomes show some similar characteristics such as genome compaction [22], AT bias [7, 36, 37], codon bias [38, 39] and extreme divergence. A summary of the features of organelle-, obligate-intracellular- and nucleomorph-genomes is given in Table 1. These features are important to consider as measure of how unusual, or not, nucleomorph genomes are.

With the recent availability of red algal [40] and green algal [41] genomic data we are for the first time in a position to do comparative genomics between nucleomorphs of both cryptomonads and chlorarachniophytes and examples of their free-living relatives, with the plant Arabidopsis thaliana serving as an outgroup. Here we test whether the phylogenetically distinct nucleomorph genomes of G. theta and B. natans have experienced similar evolutionary pressures that influenced genome-wide variation in predictable ways and with the same severity and whether these effects are in common to those described in other enslaved nuclei. Proteins from both nucleomorph genomes have been observed to reside on long branches of phylogenetic trees indicating that they are poorly conserved [42–45], however this has never been investigated at the genomic level. It is also assumed that nucleomorph genes are highly derived because the proteins function within a sub-cellular compartment, the periplastidial space, where selection is relaxed due to reduced interactions with other proteins. However, both the G. theta and B. natans nucleomorphs encode proteins that are directed to the plastid. Proteins that function in the plastid are presumably subject to similar selection pressures in organisms with nucleomorphs as they are in other algae. We have therefore used plastid proteins encoded in the plastid genome, the nucleomorph, or the nucleus, to examine differences in rates of evolution in the different genomes to determine whether the nucleomorph is evolving at a dissimilar rate to the plastid and nuclear genomes. We also investigate the overall variability of evolutionary rates of nucleomorph-encoded proteins and their homologues in other species to determine if the proteins still encoded within these genomes are generally well conserved, and whether this can shed light on their retention in the nucleomorph. By comparing proteins from the nucleomorph of two cryptomonads, G. theta and Rhodomonas salina, we also investigate whether cryptomonad nucleomorph genomes are diverging at the same rate as their nuclear genomes.


Mitochondria

Mitochondria (singular = mitochondrion) are often called the “powerhouses” or “energy factories” of a cell because they are responsible for making adenosine triphosphate (ATP), the cell’s main energy-carrying molecule. The formation of ATP from the breakdown of glucose is known as cellular respiration. Mitochondria are oval-shaped, double-membrane organelles (Figure 1) that have their own ribosomes and DNA. Each membrane is a phospholipid bilayer embedded with proteins. The inner layer has folds called cristae, which increase the surface area of the inner membrane. The area surrounded by the folds is called the mitochondrial matrix. The cristae and the matrix have different roles in cellular respiration.

In keeping with our theme of form following function, it is important to point out that muscle cells have a very high concentration of mitochondria because muscle cells need a lot of energy to contract.

Figure 1 This transmission electron micrograph shows a mitochondrion as viewed with an electron microscope. Notice the inner and outer membranes, the cristae, and the mitochondrial matrix. (credit: modification of work by Matthew Britton scale-bar data from Matt Russell)

Like mitochondria, chloroplasts also have their own DNA and ribosomes. Chloroplasts function in photosynthesis and can be found in eukaryotic cells such as plants and algae. Carbon dioxide (CO2), water, and light energy are used to make glucose and oxygen in photosynthesis. This is the major difference between plants and animals: Plants (autotrophs) are able to make their own food, like glucose, whereas animals (heterotrophs) must rely on other organisms for their organic compounds or food source.

Like mitochondria, chloroplasts have outer and inner membranes, but within the space enclosed by a chloroplast’s inner membrane is a set of interconnected and stacked, fluid-filled membrane sacs called thylakoids (Figure 2). Each stack of thylakoids is called a granum (plural = grana). The fluid enclosed by the inner membrane and surrounding the grana is called the stroma.

Figure 2 This simplified diagram of a chloroplast shows the outer membrane, inner membrane, thylakoids, grana, and stroma.

The chloroplasts contain a green pigment called chlorophyll, which captures the energy of sunlight for photosynthesis. Like plant cells, photosynthetic protists also have chloroplasts. Some bacteria also perform photosynthesis, but they do not have chloroplasts. Their photosynthetic pigments are located in the thylakoid membrane within the cell itself.

Theory of Endosymbiosis

We have mentioned that both mitochondria and chloroplasts contain DNA and ribosomes. Have you wondered why? Strong evidence points to endosymbiosis as the explanation.

Symbiosis is a relationship in which organisms from two separate species live in close association and typically exhibit specific adaptations to each other. Endosymbiosis (endo-= within) is a relationship in which one organism lives inside the other. Endosymbiotic relationships abound in nature. Microbes that produce vitamin K live inside the human gut. This relationship is beneficial for us because we are unable to synthesize vitamin K. It is also beneficial for the microbes because they are protected from other organisms and are provided a stable habitat and abundant food by living within the large intestine.

Scientists have long noticed that bacteria, mitochondria, and chloroplasts are similar in size. We also know that mitochondria and chloroplasts have DNA and ribosomes, just as bacteria do. Scientists believe that host cells and bacteria formed a mutually beneficial endosymbiotic relationship when the host cells ingested aerobic bacteria and cyanobacteria but did not destroy them. Through evolution, these ingested bacteria became more specialized in their functions, with the aerobic bacteria becoming mitochondria and the photosynthetic bacteria becoming chloroplasts.