Content uploaded by Elvin J Lauron
Author content
All content in this area was uploaded by Elvin J Lauron on Oct 21, 2014
Content may be subject to copyright.
Available via license: CC BY 4.0
Content may be subject to copyright.
R E S E A R C H Open Access
Transcriptome sequencing and analysis of
Plasmodium gallinaceum reveals polymorphisms
and selection on the apical membrane antigen-1
Elvin J Lauron
1*
, Khouanchy S Oakgrove
1
, Lisa A Tell
2
, Kevin Biskar
1
, Scott W Roy
1
and Ravinder NM Sehgal
1
Abstract
Background: Plasmodium erythrocyte invasion genes play a key role in malaria parasite transmission, host-specificity
and immuno-evasion. However, the evolution of the genes responsible remains understudied. Investigating these
genes in avian malaria parasites, where diversity is particularly high, offers new insights into the processes that confer
malaria pathogenesis. These parasites can pose a significant threat to birds and since birds play crucial ecological roles
they serve as important models for disease dynamics. Comprehensive knowledge of the genetic factors involved in
avian malaria parasite invasion is lacking and has been hampered by difficulties in obtaining nuclear data from avian
malaria parasites. Thus the first Illumina-based de novo transcriptome sequencing and analysis of the chicken parasite
Plasmodium gallinaceum was performed to assess the evolution of essential Plasmodium genes.
Methods: White leghorn chickens were inoculated intravenously with erythrocytes containing P. gallinaceum.cDNA
libraries were prepared from RNA extracts collected from infected chick blood and sequencing was run on the
HiSeq2000 platform. Orthologues identified by transcriptome sequencing were characterized using phylogenetic, ab
initio protein modelling and comparative and population-based methods.
Results: Analysis of the transcriptome identified several orthologues required for intra-erythrocytic survival and
erythrocyte invasion, including the rhoptry neck protein 2 (RON2) and the apical membrane antigen-1 (AMA-1).
Ama-1of avian malaria parasites exhibits high levels of genetic diversity and evolves under positive diversifying
selection, ostensibly due to protective host immune responses.
Conclusion: Erythrocyte invasion by Plasmodium parasites require AMA-1 and RON2 interactions. AMA-1 and
RON2 of P. gallinaceum are evolutionarily and structurally conserved, suggesting that these proteins may play essential
roles for avian malaria parasites to invade host erythrocytes. In addition, host-driven selection presumably results in the
high levels of genetic variation found in ama-1 of avian Plasmodium species. These findings have implications for
investigating avian malaria epidemiology and population dynamics. Moreover, this work highlights the P. gallinaceum
transcriptome as an important public resource for investigating the diversity and evolution of essential Plasmodium
genes.
Keywords: Avian malaria, Transcriptome, Apical membrane antigen I, Synonymous, Non-synonymous, Polymorphism
* Correspondence: elauron@mail.sfsu.edu
1
Department of Biology, San Francisco State University, San Francisco, CA
94132, USA
Full list of author information is available at the end of the article
© 2014 Lauron et al.; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative
Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and
reproduction in any medium, provided the original work is properly credited. The Creative Commons Public Domain
Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article,
unless otherwise stated.
Lauron et al. Malaria Journal 2014, 13:382
http://www.malariajournal.com/content/13/1/382
Background
Immuno-evasion is likely a major factor that influences
the evolution of Plasmodium parasites, the causative
agent of malaria [1]. Parasites with antigenic diversity
may have frequency-dependent advantages and as a re-
sult evolve under strong positive/diversifying selection
[2]. The genetic diversity maintained by positive selec-
tion in target malaria antigens poses a major problem in
the development of effective malaria vaccines [3,4]. In
general, most genes that encode antigens of Plasmodium
parasites are highly polymorphic and encode proteins
that are important targets for host protective antibody
responses [5]. Indeed, polymorphisms in the circumspor-
ozoite protein, a cell surface protein required for sporo-
zoites to attach and invade target cells, appear to be
maintained by selective pressures exerted via host pro-
tective immune responses [6,7]. Evidence for positive se-
lection has also been reported for the Plasmodium
surface proteins DBP (Duffy-binding protein), EBA-175
(erythrocyte-binding antigen 175) and a large number of
other antigens [8].
Polymorphisms maintained by selective pressures within
erythrocyte-binding ligands may alter the host receptor-
specificity of Plasmodium parasites [3,9,10]. In Plasmo-
dium parasites that infect multiple host species through
host-switching events, e.g., avian malaria species, such
polymorphisms may contribute to a broad host-specificity
range [11]. The host-specificity of avian malaria parasites
is diverse: some parasites can infect hosts from multiple
families or even multiple orders; others are restricted to
a single avian family or even species [12-14]. Therefore,
avian Plasmodium parasites provide an exceptional
model for studying host specificity and host-parasite
co-evolutionary dynamics in natural populations [15,16].
Host-parasite co-evolutionary relationships are thought
to maintain genetic diversity in both host and parasite
populations [17]. Indeed, there is evidence for parasite-
driven diversifying selection in avian hosts [18-20]. How-
ever, little is known regarding host-driven selection in
avian Plasmodium parasites. This is largely due to the dif-
ficulties of identifying and obtaining data on nuclear
genes: since erythrocytes are nucleated in bird hosts, it
is hard to isolate parasite DNA/RNA from the much
more abundant host material [21]. Therefore, Plasmo-
dium gallinaceum, a parasite of the domestic chicken
(Gallus gallus), was chosen to perform transcriptome
sequencing and analysis in this study since it is rela-
tively easy to propagate in chickens and generating high
parasitemia is readily achieved with this strain.
Plasmodium gallinaceum has been an important model
for understanding cellular biological mechanisms involved
in malaria parasite transmission [22-24], and can yield
insight applicable to Plasmodium falciparum [25], as
P. falciparum shares high similarity with the genome
of P. gallinaceum. This is supported by phylogenetic
evidence [26-31] and biochemical data that function-
ally confirm the evolutionary relationships [25]. Here,
the goal was to identify orthologues of essential and
well-characterized P. falciparum genes from the P. gal-
linaceum transcriptome; some of these include the
long chain fatty acid elongation enzyme (ELO3), LCCL
domain-containing protein (CCp2), and serine hydro-
xymethyltransferase (SHMT).
SHMT is highly upregulated throughout the intra-
erythrocytic development stages [32] and plays an indis-
pensable role in the de novo pyrimidine biosynthesis
pathway in Plasmodium parasites; the essentiality of
SHMT has been confirmed through SHMT-knockout
parasites [33]. ELO3 and CCp2 also play essential roles
during intra-erythrocytic development and are specific-
ally required for Plasmodium gametocytogenesis, as
demonstrated by transposon-mediated insertional muta-
genesis [34]. In addition, two orthologues that are essen-
tial for erythrocytic invasion, AMA-1 (a major malaria
vaccine candidate) and RON2 (the AMA-1 receptor)
were characterized in this study.
Plasmodium invasion of erythrocytes can be blocked
by antibody-mediated inhibition of AMA-1-RON2 inter-
actions [35-37], and the vaccine potential of AMA-1 has
been well demonstrated in various animal models
[38-40]. In spite of these promising results, different iso-
lates of the same species exhibit polymorphisms in ama-
1that may allow parasites to avoid inhibitory effects of
natural anti-AMA1 antibodies produced by host protect-
ive immune responses [41]. Moreover, natural immune
responses to AMA-1 have also revealed polymorphic B
and T cell epitopes within AMA-1 that are maintained
by positive selection [42,43]. Given that AMA-1 is a
highly polymorphic antigen that is unique to apicom-
plexan parasites [44,45], the level of diversity and selec-
tion on avian Plasmodium ama-1was evaluated. The
results of this study have implications for studying
erythrocyte invasion, host immune responses and the
population genetics and epidemiology of avian malaria
parasites.
Methods
Infection of chickens
A total of seven White Leghorn chickens were hatched
at the animal facility of the University of California,
Davis, and were kept in cages with water and feed. After
six days, six chicks were inoculated intravenously with
erythrocytes containing P. gallinaceum. One chick was
inoculated with saline solution as a negative control.
Blood samples were obtained from the jugular vein
seven days post infection and subsequently every two to
three days. Blood samples were stored in TRIzol® RNA
Isolation Reagent (Life Technologies, USA) and were
Lauron et al. Malaria Journal 2014, 13:382 Page 2 of 14
http://www.malariajournal.com/content/13/1/382
immediately flash frozen in an ethanol-dry ice bath, or
used for blood smear examination. Blood smears were
stained with Giemsa, and the infection status was veri-
fied by microscopy and PCR amplification of the cyto-
chrome b gene [46].
Generation of cDNA libraries, sequencing and data
analysis of the Plasmodium gallinaceum blood stages
Total RNA was prepared directly from the frozen sam-
ples of parasitized erythrocytes. RNA was extracted
using Phase Lock Gel and ethanol precipitation methods
[47]. The RNA quality was checked on the Bioanalyzer
2100 (Agilent Technologies Inc., USA). cDNA libraries
were prepared from RNA extracts and sequenced at the
qb3 Genomics Sequencing Laboratory at the University
of California, Berkeley, USA as follows: rRNA was de-
pleted from RNA extracts using Ribo-Zero™(Epicentre,
USA) prior to generating cDNA libraries using TruSeq™
(Illumina Inc, USA). Sequencing was run in one lane as
paired-end reads of 100 base pairs (bp) on the HiSeq2000
platform. The quality of all Illumina reads was assessed
with FastQC [48]. Overall, the sequence reads were of
good quality (average quality score of 38 per read).
Seventy-five per cent of the sequence reads had a quality
score ≥30. Thus, no quality trimming was required nor
performed, so as to minimize loss of the dataset. Blat/
Bowtie [49,50] query of the Illumina reads against the
G. gallus (chicken) genome was run to remove chicken
sequences. Adapters were removed and the remaining
paired-end reads were used for the de novo reconstruc-
tion of the P. gallinaceum transcriptome using Trinity
[51]. To identify P. gallinaceum protein-coding transcripts
involved in erythrocyte invasion, intra-erythrocytic sur-
vival and gametocytogenesis, Tblastn of the P. gallinaceum
transcriptome against the P. falciparum transcriptome
was performed using Geneious 7.0.4. An E-value cut-off of
1e-10 was chosen for identifying putative orthologues.
The P. falciparum transcriptome was downloaded from
PlasmoDB [52].
Sample collection
Plasmodium lucens isolates used in this study came from
blood samples collected from a single species, the Olive
Sunbird (Cyanomitra olivacea), in Cameroon during the
period 2005 to 2007 [46]. Plasmodium globularis was
isolated from blood samples collected from the Yellow-
whiskered Greenbul (Andropadus latirostris) in Ghana,
2007 [53]. Plasmodium megaglobularis isolates came
from blood samples collected from the Olive-bellied
Sunbird (Cinnyris chloropygius) in Cameroon during
1990 [53]. Plasmodium lineage spp. PV16 isolates were
from blood samples collected from the Olive Sunbird in
Cameroon during 2005 [14]. Plasmodium homopolare
isolates were collected from various birds in China Creek
County Park, California, USA (Additional file 1) during
2011 to 2013 [54]. All birds were caught with mist-nets
and banded. Blood samples were collected from the bra-
chial vein and samples were stored in lysis buffer (10 mM
Tris-HCL pH 8.0, 100 mM EDTA, 2% SDS).
PCR amplification and DNA sequencing
DNA was extracted from whole blood following a
DNeasy kit protocol (Qiagen, USA). Identification of
avian Plasmodium species was based on PCR assays and
sequences of the cytochrome b gene [55]. The P. gallina-
ceum ama-1 coding sequence was identified using the P.
gallinaceum RNA-seq data, and ama-1 domain I
primers were designed based on conserved regions
among P. gallinaceum and other mammalian Plasmo-
dium species. A nested PCR was used to amplify the hy-
pervariable domain I region of ama-1 corresponding to
444-906 bp or 271-732 bp according to P. falciparum
ama-1or P. gallinaceum ama-1, respectively. The fol-
lowing primers were used for the first round of amplifi-
cation: Pg_AMA1F1 (GATTTAGGTGAAGATGCAGAA
GT) and Pg_AMA1R1 (TTAATTAAACATGTTGGTTT
TACAT). The amplification conditions were as follows,
first, 4 min at 94°C, followed by 20 cycles with 0.5 min
of denaturation at 94°C, annealing at 50°C for 1 min,
and elongation at 72°C for 1.2 min. After 20 cycles, a
final elongation step at 72°C for 5 min was carried out.
The amplified products of 785 bp were used for the sec-
ond round of amplification with the following primers:
Pg_AMA1F2 (ATGTCCAGTTTTTGGAAAAGGTAT) and
Pg_AMA1R2 (CCATCAACCCATAAT CCAAATTT). The
second round amplification conditions were as follows first,
1minat94°C,followedby40cycleswith0.5minofde-
naturation at 94°C, annealing at 53°C for 1 min, and elong-
ation at 72°C for 0.7 min. After 40 cycles, a final elongation
step at 72°C for 5 min was carried out. The amplified prod-
ucts of 500 bp were run out on a 1.8% agarose gel using 1 ×
TBE, and visualized by ethidium bromide staining under
ultraviolet light. Resulting amplicons were purified using
ExoSap (following manufacture’s instructions, USB Corp,
USA) and sequenced by ElimBio (Hayward, USA), see
Additional file 2 and Additional file 3 for accession num-
bers. Several attempts to amplify domain II and III were
unsuccessful, which may have been due to the low GC
content in these regions and the difficulty in designing
highly specific primers.
Phylogenetic analyses
DNA sequences were analysed using Geneious v7.0.4
created by Biomatters. Sequences were aligned using
MUSCLE with Seaview software [56]. DNA sequences
were translated and adjusted in Mesquite v2.75 [57].
Phylogenetic relationships were inferred using maximum
likelihood (ML), as implemented in RAxML [58]. For
Lauron et al. Malaria Journal 2014, 13:382 Page 3 of 14
http://www.malariajournal.com/content/13/1/382
ML, Modeltest v3.7 [59] was used to determine the most
appropriate nucleotide substitution model based on the
Akaike Information Criterion (AIC) [60]. ML methods
for the ama-1and SHMT genes were implemented using
the GTR + I + G model that permits rate variations in all
six base substitution types for unequal base composition,
invariable sites and among site rate variation. ML
methods for the RON2,ELO3,CCp2 genes were imple-
mented using the GTR + G model. A thorough ML search
was performed along with 10,000 bootstrap inferences.
Protein structure modelling
Three-dimensional (3D) models representing tertiary
protein structures of AMA-1, domain I of AMA-1 alone,
and RON2 was generated using an ab initio approach
with the iterative implementation of the threading as-
sembly refinement (I-TASSER) method [61,62]. The ac-
curacy of the 3D models was assessed based on the
confidence (C) score and the template modelling (TM)
score. The quality of the top-ranked 3D models, mea-
sured by LGscore and MaxSub values, was further
assessed using protein quality predictor ProQ [63]. 3D
models with an LGscore greater than 2.5 are considered
very good models, where as values above 4 indicate ex-
tremely good models. 3D models with MaxSub values
above 0.1 are considered fairly good models, whereas
values above 0.5 indicate very good models. Secondary
structures of AMA-1 or domain I of AMA-1 alone and
RON2 were determined by PSIPRED v3.3 [64].
Statistical analyses of genetic diversity
A total of 51 P. lucens ama-1sequences consisting of 12
different haplotypes were compared with 49 P. falcip-
arum ama-1sequences. The McDonald-Kreitman test
[65] was performed on domain I of ama-1 to determine
whether this region is evolving under selection. The d
N
/
d
S
(non-synonymous substitutions per non-synonymous
sites divided by synonymous substitutions per synonym-
ous sites) ratio was evaluated using a sliding window
method to investigate selection across the region, as im-
plemented in DNAsp v5.10 [66]. Significant differences
between d
N
and d
S
were evaluated for the entire region
of domain I and for regions with high d
N
/d
S
ratios using
the Nei and Gojobori method with the Jukes and Cantor
correction, and a one-tailed Z-test with 1,000 bootstrap
pseudosamples, as implemented in MEGA v5.2.2 [67].
Between-species divergence (K) using Jukes and Cantor
correction was calculated with MEGA (see Additional
file 4 and Additional file 5). The nucleotide diversity (π)
across domain I of P. lucens ama-1 was also evaluated
using a sliding window method.
Tajima’s test was performed to determine if sequences
departed significantly from neutral variation patterns.
With Tajima’s test, departure from neutrality is measured
by differences between πand the nucleotide diversity ex-
pected under neutrality (θ). πis expected to increase
above that of θas a result of a rare allele being selected
and maintained at intermediate frequencies under posi-
tive selection. Thus, a positive test statistic (D) value
under positive diversifying selection is expected [68,69].
Fu and Li’s test was also performed using P. gallinaceum
as outgroup to determine whether mutations are select-
ively neutral. Similarly, a positive value of D* and F*
under positive diversifying selection is expected. When
comparing estimates of θbased on singleton sites to that
derived from the D*orF* index, an excess of intermediate
frequency polymorphisms and lower number of singleton
sites makes the statistics values positive [69,70]. To check
for clustering, a metric multidimensional scaling analysis
was performed in R using the bios2mds package.
Results
Transcriptome sequencing
A total of 100 Gb comprising of 220 M 100 nucleotide
(nt) paired-end sequencing read was obtained. Sixty-
three percent of the total sequence reads obtained were
removed after running a Bowtie query against the G.
gallus genome. The remaining 82 M sequence-read pairs
were assembled de novo using Trinity. Long open read-
ing frames (ORFs) within the assembled transcriptome
were identified. These putative coding sequences (CDS)
were compared to the P. falciparum (isolate 3D7) tran-
scriptome. Eighty-one per cent of P. falciparum CDS
sequences had a significant BLAST hit within the P. gal-
linaceum transcriptome, suggesting that the P. gallina-
ceum transcriptome obtained was fairly complete. The
size distribution for the CDS that showed homology to
P. falciparum CDS is shown in Additional file 6.
Identification and phylogenetic analysis of genes
essential for intra-erythrocytic stage survival, and
gametocytogenesis
A full-length cDNA sequence encoding an orthologue of
SHMT in P. gallinaceum, with an 84% amino acid se-
quence identity in a pairwise comparison to P. falciparum
SHMT, was identified.The translated SHMT sequences of
all analysed Plasmodium species resulted in proteins with
identical lengths of 442 amino acids. Twelve cysteines
were present in P. gallinaceum SHMT, eight of which
were conserved in position.
In addition, ELO3 and CCp2 were also identified as
orthologues in P. gallinaceum. The full-length cDNA se-
quence was obtained for ELO3 and a partial cDNA se-
quence was obtained for CCp2. The 5’exon sequence of
CCp2 was missing approximately 650 bp. The resulting
coding sequences of ELO3 and CCp2 are 1,920 and
4,201 bp, respectively. DNA sequences were translated
and aligned; the resulting amino acid sequences showed
Lauron et al. Malaria Journal 2014, 13:382 Page 4 of 14
http://www.malariajournal.com/content/13/1/382
79 and 74% identity in pairwise comparisons to P. falcip-
arum ELO3 and CCp2, respectively. The P. gallinaceum
CCp2 amino acid sequence (1,348 amino acids) that was
analysed contains 17 cysteines, all of which were conserved
between mammalian Plasmodium species. The translated
P. gallinaceum ELO3 full-length transcript sequence re-
sulted in a protein of 521 amino acids and was approxi-
mately 121 amino acids shorter than the P. falciparum
ELO3 protein. Phylogenetic analyses suggested that these
essential orthologues in P. gallinaceum are most similar
to P. falciparum (as compared to other mammalian
Plasmodium species) (Figure 1A-C). The high degree of
conservation suggests that these orthologues may also
play important roles during the intra-erythrocytic
stages of P. gallinaceum.
Identification and phylogenetic analysis of ama-1 and the
ama-1 receptor RON2 in Plasmodium gallinaceum
The coding region of P. gallinaceum ama-1was 1,692 bp
with an A + T content (72%) greater than P. falciparum
(around 70%). The translated protein sequence was 556
amino acids long, shared 55% amino acid identity to P. fal-
ciparum ama-1, and contains 17 cysteines. All 16 cyste-
ines within the three cysteine-rich domains of AMA-1
Figure 1 Phylogeny of Plasmodium parasites based on A) 1,329 bp of the serine hydroxymethyltransferase (SHMT) gene, B) 1920 bp of
the long chain fatty acid elongation enzyme ( ELO3) gene, C) 4854 bp of the LCCL domain-containing protein (CCp2) gene, and
D) 7563 bp of the Rhoptry neck protein 2 (RON2) gene. Numbers on the branches refer to bootstrap values obtained with 10,000
replicates. For accession numbers see Additional file 2.
Lauron et al. Malaria Journal 2014, 13:382 Page 5 of 14
http://www.malariajournal.com/content/13/1/382
were conserved in number and position when the aligned
protein sequences of individual domains were analysed,
with the exception of domain III. Domain III contains six
cysteine residues [71,72], which were present in both P.
gallinaceum and P. falciparum ama-1. However, domain
III of P. gallinaceum AMA-1 contained five amino acid
deletions between positions 465-471 (relative to P. falcip-
arum AMA-1). Therefore, the position of cysteines and
the number of amino acids in domain III varied slightly
between P. gallinaceum and P. falciparum. Phylogenetic
analysis of ama-1revealed P. gallinaceum ama-1as sig-
nificantly divergent from all mammalian Plasmodium spe-
cies analysed (Figure 2). A metric multidimensional
scaling (MDS) analysis was performed to complement the
phylogeny and to visualize the evolutionary trajectories of
ama-1on a low dimensional space. Principal component
analysis (PCA) plots showed clustering consistent with
that of the ama-1phylogeny (Figure 2). Phylogenetic and
metric multidimensional scaling analysis of all ama-1se-
quences compared in this study placed avian Plasmodium
parasites into a strongly supported monophyletic clade
and cluster (Additional file 7).
In addition to identifying AMA-1 in P. gallinaceum,a
full-length cDNA sequence encoding a version of P.
gallinaceum RON2 was identified. P. gallinaceum RON2
was truncated at the N terminus by approximately 620
amino acids, in comparison to the translated full-length
cDNA sequence of P. falciparum RON2. The coding se-
quence of P. gallinaceum RON2 was 4,641 bp and en-
codes a protein of 1545 amino acids with 71% identity to
P. falciparum RON2. Two conserved cysteines that are
required for RON2 to bind the AMA-1 pocket [73] were
conserved in P. gallinaceum RON2 (Figure 3). A phylo-
genetic analyses of RON2 groups P. gallinaceum with P.
falciparum (Figure 1D).
ama-1 polymorphisms in avian Plasmodium field isolates
The entire ama-1domain I region consisting of 468 bp
was sequenced and analysed from a total of 51 P. lucens
isolates collected from African rainforest birds. A sliding
window analysis of πusing a window of 30 bp moved in
steps of nine sites reveals polymorphisms across the en-
tire region of domain I (Figure 4A). The region 787-
816 bp appears to be the most polymorphic region in
domain I of P. lucens ama-1. This region corresponds to
the naturally immunogenic T cell epitope located within
residues 259-271 of P. falciparum AMA-1, which was
also reported to be polymorphic [42]. Three-hundred
Figure 2 Phylogeny and metric multidimensional scaling of Plasmodium parasites using ama-1.A matrix of distances (p-distance) were
generated from aligned ama-1sequences. The distance matrix was used to calculate principal components and is represented by metric
multidimensional scaling (MDS). For graphical purposes, the phylogeny was superimposed onto a PCA plot to visualize sequence space.
Lauron et al. Malaria Journal 2014, 13:382 Page 6 of 14
http://www.malariajournal.com/content/13/1/382
and eighty-seven monomorphic sites and 81 poly-
morphic sites were detected in domain I of P. lucens
ama-1. Three polymorphic sites at positions 116, 138
and 205 exhibited three different nucleotides, whereas
the remaining sites had only two. A total of 77 muta-
tions were detected, 27 of which were synonymous and
50 of which were non-synonymous. πfor all 51 se-
quences analysed was 0.043237 ± 0.005210 SD, which is
more polymorphic than previously reported π(0.01361-
0.01764) for P. falciparum isolates [42,74].
An alignment of the translated sequences revealed three
additional amino acid residues (Glu-Phe-X) positioned
near hydrophobic amino acids that line the hydrophobic
trough of domain I (Additional file 8). The additional
amino acids are located between residues 184-185 relative
to P. falciparum AMA-1. Considering that it is unknown
Figure 3 Three-dimensional models of the avian Plasmodium RON2 protein. A) Stereo views of the C-terminus of P. gallinaceum RON2 (left)
and P. falciparum RON2 (right). B) Sequence alignment of P. gallinaceum and P. falciparum RON2 are shown with the secondary structure of the
corresponding amino acid regions above the alignment. Helices are colored red. The line with connecting arrows indicates disulfide bonds.
Conserved amino acids are highlighted in red.
Figure 4 Sliding window plot of A) πand B) d
N
/d
S
for domain I of Plasmodium lucens ama-1.Nucleotide positions are relative to the
P. falciparum 3D7 line sequence. The window length is 30 bp with a step size of 9 bp for the sliding window analysis of π. The window length is 90 bp
with a step size of 3 bp for the sliding window analysis of d
N
/d
S
. Asterisks indicate regions with a significant excess of non-synonymous substitutions.
Lauron et al. Malaria Journal 2014, 13:382 Page 7 of 14
http://www.malariajournal.com/content/13/1/382
whether the tertiary structure of AMA-1 in avian Plas-
modium species contains a hydrophobic trough, a 3D
model of the full-length P. gallinaceum AMA-1 was gen-
erated. Tertiary structure-based analysis of the resulting
3D model was found to be satisfactory (LGscore = 3, Max-
Sub = 0.3). Conserved hydrophobic amino acid residues
that line the hydrophobic trough, according to P. falcip-
arum AMA-1 positions found by Bai et al. [75], were
highlighted in green. All highlighted residues (111, 125,
132, 144, 299, 313 relative to P. gallinaceum AMA-1) also
appeared to reside in a small yet extended pocket of P.
gallinaceum AMA-1 (Figure 5A), with the exception of
residues 193 and 194. 3D models for the AMA-1 domain
IofP. lucens and P. falciparum were also generated to
visually compare intraspecific polymorphisms (Figure 5B).
To determine whether these polymorphisms and
additional amino acids are present among other avian
Plasmodium species, domain I of ama-1from 28 P.
homopolare (a newly identified host-generalist), three
P. megaglobularis (host-generalist), three Plasmodium
lineage spp. PV16, and two P. globularis (host-specialist)
isolates [14,54] were sequenced. Between-species diver-
gence (K) ranged 0.062-0.349 (Additional file 4), approxi-
mately two to three-fold greater than K calculated for
cytochrome b (0.030-0.103). Attempts to sequence domain
I from other avian species, including Plasmodium relictum
(GRW11 and SGS1) isolates were unsuccessful, likely due
to low parasitaemia or parasite genomic DNA concentra-
tions. Interestingly, no intraspecific polymorphisms were
observed. However, additional amino acids in domain I
Figure 5 Three-dimensional models of the avian Plasmodium AMA-1 protein. The ab initio-generated models are based on the A) 556 aa
sequence of P. gallinaceum AMA-1, B) 155 aa residues in AMA-1 domain I of P. lucens, and C) 152 aa residues in AMA-1 domain I of P. falciparum.
Both stereo (left) and surface (right) views are shown. Domain I, II and III of P. gallinaceum AMA-1 are coloured yellow, red and blue, respectively.
Conserved hydrophobic residues that line the putative hydrophobic trough are labelled and highlighted in green. Plasmodium lucens and P. falciparum
AMA-1 models are coloured as follows: conserved hydrophobic residues are shown in blue, highly polymorphic residues are shown in red, high-frequency
dimorphisms are shown in pink, low-frequency dimorphisms are shown in orange. The observed amino acid insertions in domain I of P. lucens AMA-1 are
shown in yellow.
Lauron et al. Malaria Journal 2014, 13:382 Page 8 of 14
http://www.malariajournal.com/content/13/1/382
were present in all field-caught avian Plasmodium spe-
cies included in this study. Similar to P. lucens,domain
IofP. megaglobularis, lineage PV16, and P. globularis
AMA-1 sequences contain three additional amino acids
(Glu-Phe-X) between residues 184-185, whereas the do-
main I of P. homopolare AMA-1 contains two amino
acid (Arg-Asp) insertions between residues 187-188
(Additional file 8). All additional amino acids observed
were located within or near hydrophobic amino acids
that line the hydrophobic pocket.
Comparison of Plasmodium lucens ama-1 and Plasmodium
falciparum ama-1 sequences
Forty-nine P. falciparum ama-1sequences were com-
pared with 51 P. lucens ama-1sequences in an align-
ment covering the entire domain I region. There were a
total of 120 fixed nucleotide differences between the spe-
cies. Of these differences, 33% (39) were synonymous
and 67% were (81) non-synonymous. There were a total
of 121 polymorphic sites within-species, of which 22%
(27) sites were synonymous and 78% (94) were non-
synonymous. A McDonald-Kreitman test with domain I
sequences detected significant departure from neutrality
in the P. lucens ama-1domain I region (Neutrality Index
(NI) = 1.676, P = 0.07; NI with Jukes and Cantor correc-
tion = 2.035, P = 0.009), suggesting that polymorphisms
at the domain I are maintained under positive diversify-
ing selection.
Ad
N
/d
S
analysis of the entire domain did not detect
significant differences between non-synonymous and
synonymous changes. To determine if regions with high
d
N
/d
S
ratios are under selection, a sliding-window ana-
lysis (90 bp with a step size of three bases) of d
N
/d
S
was
conducted. Significant difference between d
N
and d
S
was
detected throughout domain I (at the midpoint nt position
of region 543-650, d
N
/d
S
= 3.51, P < 0.0005; mid point nt
position of region 609-698, d
N
/d
S
= 1.90, P < 0.05; mid
point nt position of region 645-735, d
N
/d
S
= 1.97, P <
0.05; mid point nt position of region 657-746, d
N
/d
S
=
1.66, P = 0.05; mid point nt position of region 669-758,
d
N
/d
S
= 2.01, P < 0.05) (Figure 4B), suggesting that these
regions are under positive diversifying selection.
No significant departure from the neutral expectation
was detected using Tajima’sD(D= -0.12236, P > 0.10).
Similar results were obtained for the Fu and Li D*
(1.32144, P > 0.10), whereas the F* value of 1.79767 was
significant with P < 0.02. Sliding-window analysis with a
window of 30 bp and a step size of nine bases did not
detect significantly low or high statistics values for all
neutrality tests along the domain I region of ama-1.To
ensure that the true level of variation is not obscured
from choosing too small of a window, the sliding-
window analysis with a window of 90 bp and a step size
of three bases was performed. Significantly high F*
values were observed between 658-786 bp and 718-
831 bp (Figure 6). These results of the population-based
methods also support the findings from comparing d
N
to d
S
, and provide additional evidence that the domain I
region of P. lucens ama-1is under positive diversifying
selection.
Discussion
Assessing the genetic diversity and selection in erythro-
cyte invasion genes of avian malaria parasites may pro-
vide insight on the population and the transmission
dynamics of malaria. Higher rates of malaria transmis-
sion are expected to occur in areas with high parasite di-
versity [76]. This prediction is supported by the high levels
of diversity in nuclear genes from malaria parasites found
in avian hosts of Hawaii, including the thrombospondin-
related anonymous protein (trap) gene, which encodes a
protein involved in immuno-evasion and erythrocyte inva-
sion [77]. Although trap evolves under positive selection
in human malaria parasites, no evidence for positive selec-
tion was found in trap of avian malaria parasites [78].
Here, for the first time, evidence of positive selection as a
driving force in the evolution and diversification of an
erythrocyte invasion gene (ama-1) in avian malaria para-
sites is provided. These results show that ama-1is a useful
nuclear marker for investigating the adaptive evolution of
avian malaria parasite populations. In addition, these re-
sults suggest that the role ama-1plays in erythrocyte inva-
sion is evolutionarily conserved across avian Plasmodium
species.
Avian Plasmodium ama-1is relatively conserved in
comparison with orthologous genes of mammalian Plas-
modium species. The conservation of the hydrophobic
trough in P. gallinaceum AMA1 underscores the func-
tional importance of AMA-1 in avian Plasmodium para-
sites. The hydrophobic trough of AMA-1 binds to the
RON complex via AMA-1-RON2 interactions [73,79].
These interactions form the invasion machinery required
to mediate erythrocyte invasion [80,81]. Moreover, the
identification of P. gallinaceum RON2 and the conserva-
tion of the RON2 helices involved in AMA-1 binding
further supports the notion that this junction-dependent
invasion process is evolutionarily conserved in avian
Plasmodium parasites. Future studies will be important
to investigate the signatures of selection within regions
of P. gallinaceum RON2, and to further the understand-
ing of junction-dependent invasion processes in avian
Plasmodium parasites.
Although this process is not specific to a particular
host cell type, the binding of domain III to the erythro-
cyte membrane protein Kx suggests that AMA-1 may be
involved in host-specificity [82]. Interestingly, domain III
is not well conserved in other genera belonging to Api-
complexa [82], suggesting that the functions of this
Lauron et al. Malaria Journal 2014, 13:382 Page 9 of 14
http://www.malariajournal.com/content/13/1/382
domain are unique to Plasmodium. More importantly,
domain III of P. falciparum AMA-1 is antigenic and
elicits growth-inhibiting antibodies [83]. Domain III con-
tains two conserved immunodominant epitopes. The
first conserved epitope is located at position 459-464,
whereas the second is located at position 467-475. Do-
main III of P. gallinaceum AMA-1 lacks four amino acid
residues within the second immunodominant epitope
(between position 468-473 with respect to P. falciparum
AMA-1). These differences may also be present in other
avian Plasmodium species and possibly result in a lack
of or escape from inhibitory antibodies directed against
the second immunodominant epitope. It is thus tempting
to speculate that the observed differences in the domain
III region of P. gallinaceum AMA-1 may contribute to the
diverse host range of avian Plasmodium parasites. Unfor-
tunately, efforts to PCR amplify and assess the domain III
regions in ama-1of other avian Plasmodium parasites
were unsuccessful.
In spite of this, several avian Plasmodium species ex-
hibited genetic diversity and contained additional amino
acids present within the domain I region of AMA-1.
Figure 6 Sliding window plot of Tajima’sD, Fu and Li’sD* and F* tests for domain I of Plasmodium lucens ama-1.The window length is
90 bp with a step size of 3 bp. Asterisks indicate regions where significant departure from neutrality was observed. n represents the number of
P. lucens sequences analysed.
Lauron et al. Malaria Journal 2014, 13:382 Page 10 of 14
http://www.malariajournal.com/content/13/1/382
Plasmodium isolates from African birds contained three
additional amino acids in domain I, whereas P. homopo-
lare isolates from California contained two additional
amino acids. Surprisingly, domain I of P. homopolare
ama-1was highly conserved with no genetic diversity
found between 28 different isolates. One possibility for
this finding is that domain I of P. homopolare ama-1is
subject to negative selection, as negative selection has
been shown on the rhoptry-associated protein 1 (RAP-1),
which is also involved in erythrocyte invasion [84]. How-
ever, this possibility is difficult to investigate considering
the lack of diversity in P. homopolare ama-1.Alterna-
tively, the lack of diversity may result from a recent demo-
graphic sweep (bottle-neck) or host switching events that
lead to recent population expansions [85,86]. The latter is
favoured, as P. homopolare was found in five different
families representing nine bird species [54]. Therefore,
any reciprocal selection acting on P. homopolare ama-1
by host protective immune responses may be weak rela-
tive to the selection imposed on a host specialist by a sin-
gle host (e.g., the Olive Sunbird and P. lucens). A lack of
diversity was also found in the host generalist P. megaglo-
bularis. However, more robust sampling and sequencing
ofparasitesisrequiredasthetruediversitymaybe
underestimated due to low sample sizes. These results
are in stark contrast to the level of genetic diversity
found in P. lucens ama-1.
These findings provide significant evidence that poly-
morphisms in P. lucens ama-1are maintained by posi-
tive selection. The patterns of genetic diversity across
domain I of P. lucens ama-1are consistent with studies
on mammalian Plasmodium species [72,74,87,88]. Like-
wise, sliding window plots of πand Fu and Li’sF* value
indicate that the region corresponding to a natural T cell
epitope in domain I is highly polymorphic and is under
positive selection in P. lucens ama-1, which is also in
agreement with earlier studies [42,89]. This observation
suggests that avian Plasmodium AMA-1 may induce T
cell responses in infected bird hosts and has implications
for studying immune responses in bird populations that
are naturally exposed to malaria parasites. In addition,
humoral immune responses against AMA-1 in bird pop-
ulations may also provide valuable information for
immuno-epidemiologic studies. Such studies can be par-
ticularly important for terrestrial ecosystems that are sen-
sitive to losses in native bird populations, especially since
native birds play essential roles in many terrestrial ecosys-
tems [90,91]. Therefore, there is a great need to under-
stand host-parasite interaction and its relationship to host
immune responses in avian Plasmodium parasites.
Conclusions
Ultimately, these findings provide insight into the
erythrocyte invasion process of avian Plasmodium, are
the first evidence of host-driven selection in an avian
Plasmodium species, and demonstrate the substantial
applications of the P. gallinaceum transcriptome. The P.
gallinaceum transcriptome dataset represents a major
public genomic resource that will serve to progress re-
search on the functional genomics and evolution of Plas-
modium. Further analyses of invasion and immuno-
evasion-related genes could reveal additional nuclear
markers for phylogenetic applications, as these genes
may exhibit high levels of diversity [21]. Identifying add-
itional markers has been difficult as the majority of se-
quenced Plasmodium genomes are from mammalian
Plasmodium species; therefore, primer development is
often facilitated using mammalian Plasmodium species
sequence data and the resulting primers may not be suit-
able for PCR amplification of non-mammalian Plasmo-
dium species DNA. However, with the recent sequencing
and availability of the P. relictum transcriptome [21] and
the addition of the P. gallinaceum transcriptome, signifi-
cant progress towards adding reliable markers for more
thorough phylogenetic and evolutionary studies of Plas-
modium or closely related genera is expected.
Additional files
Additional file 1: Host and parasite species locality. The table shows
the source of isolates used to analyse sequence diversity.
Additional file 2: GenBank Accession numbers for the parasite taxa.
GenBank Accession numbers for the parasite taxa used in this study are
provided.
Additional file 3: Genbank accession numbers for the ama-1
sequences. The table shows Genbank accession numbers for the ama-1
sequences obtained from Plasmodium field isolates used in this study.
Additional file 4: Between-species divergence. The table shows
between-species divergence (K) calculated using Jukes and Cantor
correction.
Additional file 5: Genetic distances and amino acid similarity. The
table shows genetic distances computed from DNA sequences and amino
acid similarity between Plasmodium gallinaceum and Plasmodium falciparum.
Additional file 6: The size distribution for Plasmodium gallinaceum
transcripts. The figure shows the size distribution for Plasmodium
gallinaceum transcripts that show homology to Plasmodium falciparum
transcripts.
Additional file 7: Phylogenetic and MDS analysis of Plasmodium
parasites. An ama-1 phylogeny and MDS analysis shows clustering of
avian malaria parasites according to host species.
Additional file 8: AMA1- domain I amino acid sequence alignment.
The figure shows aligned amino acid sequences with the AMA-1 domain
IofPlasmodium falciparum,Plasmodium lucens,Plasmodium megaglobularis,
Plasmodium globularis,Plasmodium lineage spp. PV16, and Plasmodium
homopolare.
Competing interests
The authors declare that they have no competing interests.
Authors’contributions
EJL performed the RNA preparations, transcriptome analysis, ama-1 sequencing
and orthologue characterization, statistical analyses, and participated in the
chicken infections, blood sample collection, design of the study, and drafting of
the manuscript. KSO participated in the chicken infections, blood sample
Lauron et al. Malaria Journal 2014, 13:382 Page 11 of 14
http://www.malariajournal.com/content/13/1/382
collection and RNA preparations. LAT performed the chicken infections and
assisted with the blood sample collections. KB performed the transcriptome
assembly. SWR performed the characterization of the transcriptome. RNMS
conceived and participated in the design of the study, and helped draft the
manuscript. All authors read and approved the final version of the manuscript.
Acknowledgements
This work was supported by a National Institute of Health grant
SC2AI089120-01A1, and used the Vincent J. Coates Genomics Sequencing
Laboratory at UC Berkeley, supported by NIH S10 Instrumentation Grants
S10RR029668 and S10RR027303. We would like to thank Dr. Robert Gwadz
for kindly providing us with P. gallinaceum samples. We thank Eric Routman
(San Francisco State University) for his assistance with statistical analysis, and
thank Erika Walther and other scientists involved in sampling efforts.
Author details
1
Department of Biology, San Francisco State University, San Francisco, CA
94132, USA.
2
Department of Medicine and Epidemiology, School of
Veterinary Medicine, University of California, Davis, CA 95616, USA.
Received: 12 June 2014 Accepted: 17 September 2014
Published: 26 September 2014
References
1. Ferreira MU, da Silva Nunes M, Wunderlich G: Antigenic diversity and
immune evasion by malaria parasites. Clin Diagn Lab Immunol 2004,
11:987–995.
2. Escalante AA, Lal AA, Ayala FJ: Genetic polymorphism and natural
selection in the malaria parasite Plasmodium falciparum.Genetics 1998,
149:189–202.
3. Takala SL, Plowe CV: Genetic diversity and malaria vaccine design, testing,
and efficacy: Preventing and overcoming “vaccine resistant malaria”.
Parasite Immunol 2009, 31:560–573.
4. Roy SW, Irimia M: Origins of human malaria: Rare genomic changes and
full mitochondrial genomes confirm the relationship of Plasmodium
falciparum to other mammalian parasites but complicate the origins of
Plasmodium vivax.Mol Biol Evol 2008, 25:2511–2511.
5. Deitsch KW, Moxon ER, Wellems TE: Shared themes of antigenic variation
and virulence in bacterial, protozoal, and fungal infections. Microbiol Mol
Biol Rev 1997, 61:281–293.
6. Hughes AL: Circumsporozoite protein genes of malaria parasites
(Plasmodium spp.): evidence for positive selection on immunogenic
regions. Genetics 1991, 127:345–353.
7. Escalante AA, Grebert HM, Isea R, Goldman IF, Basco L, Magris M, Biswas S,
Kariuki S, Lal AA: A study of genetic diversity in the gene encoding the
circumsporozoite protein (CSP) of Plasmodium falciparum from different
transmission areas—XVI. Asembo Bay Cohort Project. Mol Biochem
Parasitol 2002, 125:83–90.
8. Baum J, Thomas AW, Conway DJ: Evidence for diversifying selection on
erythrocyte-binding antigens of Plasmodium falciparum and P. vivax.
Genetics 2003, 163:1327–1336.
9. Mayer DCG, Mu J-B, Feng X, Su X, Miller LH: Polymorphism in a Plasmodium
falciparum erythrocyte-binding ligand changes its receptor specificity. JExp
Med 2002, 196:1523–1528.
10. Mayer DCG, Mu J-B, Kaneko O, Duan J, Su X, Miller LH: Polymorphism in
the Plasmodium falciparum erythrocyte-binding ligand JESEBL/EBA-181
alters its receptor specificity. Proc Natl Acad Sci U S A 2004, 101:2518–2523.
11. Iyer J, Grüner AC, Rénia L, Snounou G, Preiser PR: Invasion of host cells by
malaria parasites: a tale of two protein families. Mol Microbiol 2007,
65:231–249.
12. Ricklefs RE, Fallon SM: Diversification and host switching in avian malaria
parasites. Proc R Soc Lond B 2002, 269:885–892.
13. Fallon SM, Bermingham E, Ricklefs RE: Host specialization and geographic
localization of avian malaria parasites: a regional analysis in the Lesser
Antilles. Am Nat 2005, 165:466–480.
14. Loiseau C, Harrigan RJ, Robert A, Bowie RCK, Thomassen HA, Smith TB,
Sehgal RNM: Host and habitat specialization of avian malaria in Africa.
Mol Ecol 2012, 21:431–441.
15. Bensch S, Stjernman M, Hasselquist D, Ostman O, Hansson B, Westerdahl H,
Pinheiro RT: Host specificity in avian blood parasites: a study of
Plasmodium and Haemoproteus mitochondrial DNA amplified from birds.
Proc Biol Sci 2000, 267:1583–1589.
16. Ricklefs RE, Fallon SM, Bermingham E: Evolutionary relationships,
cospeciation, and host switching in avian malaria parasites. Syst Biol
2004, 53:111–119.
17. Hellgren O, Pérez-Tris J, Bensch S: A jack-of-all-trades and still a master of
some: prevalence and host range in avian malaria and related blood
parasites. Ecology 2009, 90:2840–2849.
18. Loiseau C, Zoorob R, Robert A, Chastel O, Julliard R, Sorci G: Plasmodium
relictum infection and MHC diversity in the house sparrow (Passer
domesticus). Proc Biol Sci 2011, 278:1264–1272.
19. Sepil I, Lachish S, Hinks AE, Sheldon BC: Mhc supertypes confer both
qualitative and quantitative resistance to avian malaria infections in a
wild bird population. Proc Biol Sci 2013, 280:20130134.
20. Westerdahl H, Stjernman M, Råberg L, Lannefors M, Nilsson J-Å: MHC-I
affects infection intensity but not infection status with a frequent avian
malaria parasite in blue tits. PLoS One 2013, 8:e72647.
21. Hellgren O, Kutzer M, Bensch S, Valkiūnas G, Palinauskas V: Identification and
characterization of the merozoite surface protein 1 (msp1) gene in a host
generalist avian malaria parasite, Plasmodium relictum (lineages SGS1 and
GRW4) with the use of blood transcriptome. Malar J 2013, 12:381.
22. Carter R, Chen DH: Malaria transmission blocked by immunisation with
gametes of the malaria parasite. Nature 1976, 263:57–60.
23. Gwadz RW: Successful immunization against the sexual stages of
Plasmodium gallinaceum.Science 1976, 193:1150–1151.
24. Vinetz JM, Valenzuela JG, Specht CA, Aravind L, Langer RC, Ribeiro JMC,
Kaslow DC: Chitinases of the avian malaria parasite Plasmodium
gallinaceum, a class of enzymes necessary for parasite invasion of the
mosquito midgut. J Biol Chem 2000, 275:10331–10341.
25. Li F, Patra KP, Vinetz JM: An anti-Chitinase malaria transmission-blocking
single-chain antibody as an effector molecule for creating a Plasmodium
falciparum-refractory mosquito. J Infect Dis 2005, 192:878–887.
26. Waters AP, Higgins DG, McCutchan TF: Plasmodium falciparum appears to
have arisen as a result of lateral transfer between avian and human
hosts. Proc Natl Acad Sci U S A 1991, 88:3140–3144.
27. Escalante AA, Ayala FJ: Phylogeny of the malarial genus Plasmodium,
derived from rRNA gene sequences. Proc Natl Acad Sci U S A 1994,
91:11373–11377.
28. Escalante AA, Ayala FJ: Evolutionary origin of Plasmodium and other
Apicomplexa based on rRNA genes. Proc Natl Acad Sci U S A 1995,
92:5793–5797.
29. McCutchan TF, Kissinger JC, Touray MG, Rogers MJ, Li J, Sullivan M, Braga
EM, Krettli AU, Miller LH: Comparison of circumsporozoite proteins from
avian and mammalian malarias: biological and phylogenetic
implications. Proc Natl Acad Sci U S A 1996, 93:11889–11894.
30. Kissinger JC, Souza PCA, Soarest CO, Paul R, Wahl AM, Rathore D,
McCutchan TF, Krettli AU: Molecular phylogenetic analysis of the avian
malarial parasite Plasmodium (Novyella)juxtanucleare.J Parasitol 2002,
88:769–773.
31. Pick C, Ebersberger I, Spielmann T, Bruchhaus I, Burmester T: Phylogenomic
analyses of malaria parasites and evolution of their exported proteins.
BMC Evol Biol 2011, 11:167.
32. Nirmalan N, Wang P, Sims PFG, Hyde JE: Transcriptional analysis of genes
encoding enzymes of the folate pathway in the human malaria parasite
Plasmodium falciparum.Mol Microbiol 2002, 46:179–190.
33. Pornthanakasem W, Kongkasuriyachai D, Uthaipibull C, Yuthavong Y,
Leartsakulpanich U: Plasmodium serine hydroxymethyltransferase:
indispensability and display of distinct localization. Malar J 2012, 11:387.
34. Ikadai H, Shaw Saliba K, Kanzok SM, McLean KJ, Tanaka TQ, Cao J,
Williamson KC, Jacobs Lorena M: Transposon mutagenesis identifies
genes essential for Plasmodium falciparum gametocytogenesis. Proc Natl
Acad Sci U S A 2013, 110:E1676–E1684.
35. Deans JA, Alderson T, Thomas AW, Mitchell GH, Lennox ES, Cohen S: Rat
monoclonal antibodies which inhibit the in vitro multiplication of
Plasmodium knowlesi.Clin Exp Immunol 1982, 49:297–309.
36. Thomas AW, Deans JA, Mitchell GH, Alderson T, Cohen S: The Fab
fragments of monoclonal IgG to a merozoite surface antigen inhibit
Plasmodium knowlesi invasion of erythrocytes. Mol Biochem Parasitol 1984,
13:187–199.
37. Dutta S, Haynes JD, Moch JK, Barbosa A, Lanar DE: Invasion-inhibitory
antibodies inhibit proteolytic processing of apical membrane antigen 1
Lauron et al. Malaria Journal 2014, 13:382 Page 12 of 14
http://www.malariajournal.com/content/13/1/382
of Plasmodium falciparum merozoites. Proc Natl Acad Sci U S A 2003,
100:12295–12300.
38. Deans JA, Knight AM, Jean WC, Waters AP, Cohen S, Mitchell GH:
Vaccination trials in rhesus monkeys with a minor, invariant, Plasmodium
knowlesi 66 kD merozoite antigen. Parasite Immunol 1988, 10:535–552.
39. Collins WE, Pye D, Crewther PE, Vandenberg KL, Galland GG, Sulzer AJ,
Kemp DJ, Edwards SJ, Coppel RL, Sullivan JS: Protective immunity induced
in squirrel monkeys with recombinant apical membrane antigen-1 of
Plasmodium fragile. Am J Trop Med Hyg 1994, 51:711–719.
40. Xu H, Hodder AN, Yan H, Crewther PE, Anders RF, Good MF: CD4+ T cells
acting independently of antibody contribute to protective immunity to
Plasmodium chabaudi infection after apical membrane antigen 1
immunization. J Immunol 2000, 165:389–396.
41. Coley AM, Parisi K, Masciantonio R, Hoeck J, Casey JL, Murphy VJ, Harris KS,
Batchelor AH, Anders RF, Foley M: The most polymorphic residue on
Plasmodium falciparum apical membrane antigen 1 determines binding
of an invasion-Inhibitory antibody. Infect Immun 2006, 74:2628–2636.
42. Escalante AA, Grebert HM, Chaiyaroj SC, Magris M, Biswas S, Nahlen BL, Lal
AA: Polymorphism in the gene encoding the apical membrane antigen-1
(AMA-1) of Plasmodium falciparum. X. Asembo Bay Cohort Project. Mol
Biochem Parasitol 2001, 113:279–287.
43. Hodder AN, Crewther PE, Anders RF: Specificity of the protective antibody
response to apical membrane antigen 1. Infect Immun 2001, 69:3286–3294.
44. Michon P, Stevens JR, Kaneko O, Adams JH: Evolutionary relationships of
conserved cysteine-rich motifs in adhesive molecules of malaria
parasites. Mol Biol Evol 2002, 19:1128–1142.
45. Remarque EJ, Faber BW, Kocken CHM, Thomas AW: Apical membrane
antigen 1: a malaria vaccine candidate in review. Trends Parasitol 2008,
24:74–84.
46. Valkiūnas G, Iezhova TA, Loiseau C, Smith TB, Sehgal RNM: New malaria
parasites of the subgenus Novyella in African rainforest birds, with
remarks on their high prevalence, classification and diagnostics. Parasitol
Res 2009, 104:1061–1077.
47. Martinez C, Marzec T, Smith CD, Tell LA, Sehgal RNM: Identification and
expression of maebl, an erythrocyte-binding gene, in Plasmodium
gallinaceum.Parasitol Res 2013, 112:945–954.
48. Babraham Bioinformatics. [http://www.bioinformatics.babraham.ac.uk/]
49. Kent WJ: BLAT—The BLAST-Like Alignment Tool. Genome Res 2002,
12:656–664.
50. Langmead B, Trapnell C, Pop M, Salzberg SL: Ultrafast and memory-
efficient alignment of short DNA sequences to the human genome.
Genome Biol 2009, 10:R25.
51. Grabherr MG, Haas BJ, Yassour M, Levin JZ, Thompson DA, Amit I, Adiconis
X, Fan L, Raychowdhury R, Zeng Q, Chen Z, Mauceli E, Hacohen N, Gnirke A,
Rhind N, di Palma F, Birren BW, Nusbaum C, Lindblad-Toh K, Friedman N,
Regev A: Full-length transcriptome assembly from RNA-Seq data without
a reference genome. Nat Biotechnol 2011, 29:644–652.
52. PlasmoDB: [http://plasmodb.org]
53. Valkiūnas G, Iezhova TA, Loiseau C, Chasar A, Smith TB, Sehgal RNM: New
species of haemosporidian parasites (Haemosporida) from African
rainforest birds, with remarks on their classification. Parasitol Res 2008,
103:1213–1228.
54. Walther EL, Valkiūnas G, González AD, Matta NE, Ricklefs RE, Cornel A, Sehgal
RNM: Description, molecular characterization, and patterns of
distribution of a widespread New World avian malaria parasite
(Haemosporida: Plasmodiidae), Plasmodium (Novyella) homopolare sp.
nov. Parasitol Res 2014, 113:3319–3332.
55. Waldenström J, Bensch S, Hasselquist D, Östman Ö: A new nested
polymerase chain reaction method very efficient in detecting
Plasmodium and Haemoproteus infections from avian blood. J Parasitol
2004, 90:191–194.
56. Galtier N, Gouy M, Gautier C: SEAVIEW and PHYLO_WIN: two graphic tools
for sequence alignment and molecular phylogeny. Comput Appl Biosci
1996, 12:543–548.
57. Maddison WP, Maddison DD: Mesquite: a modular system for evolutionary
analysis. Volume Version 2.75. 2011. http://mesquiteproject.org.
58. Stamatakis A: RAxML-VI-HPC: maximum likelihood-based phylogenetic
analyses with thousands of taxa and mixed models. Bioinformatics 2006,
22:2688–2690.
59. Posada D, Crandall KA: MODELTEST: testing the model of DNA
substitution. Bioinformatics 1998, 14:817–818.
60. Huelsenbeck JP, Crandall KA: Phylogeny estimation and hypothesis
testing using maximum likelihood. Annu Rev Ecol Syst 1997, 28:437–466.
61. Zhang Y: Template-based modeling and free modeling by I-TASSER in
CASP7. Proteins 2007, 69(Suppl 8):108–117.
62. Wu S, Skolnick J, Zhang Y: Ab initio modeling of small proteins by
iterative TASSER simulations. BMC Biol 2007, 5:17.
63. Wallner B, Elofsson A: Can correct protein models be identified? Protein Sci
2003, 12:1073–1086.
64. Buchan DWA, Minneci F, Nugent TCO, Bryson K, Jones DT: Scalable web
services for the PSIPRED Protein Analysis Workbench. Nucleic Acids Res
2013, 41:W349–357.
65. Egea R, Casillas S, Barbadilla A: Standard and generalized McDonald-
Kreitman test: a website to detect selection by comparing different
classes of DNA sites. Nucleic Acids Res 2008, 36:W157–162.
66. Rozas J, Sánchez-DelBarrio JC, Messeguer X, Rozas R: DnaSP, DNA
polymorphism analyses by the coalescent and other methods.
Bioinformatics 2003, 19:2496–2497.
67. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, Kumar S: MEGA5:
molecular evolutionary genetics analysis using maximum likelihood,
evolutionary distance, and maximum parsimony methods. Mol Biol Evol
2011, 28:2731–2739.
68. Tajima F: Simple methods for testing the molecular evolutionary clock
hypothesis. Genetics 1993, 135:599–607.
69. Alexandre JS, Kaewthamasorn M, Yahata K, Nakazawa S, Kaneko O: Positive
selection on the Plasmodium falciparum clag2 gene encoding a
component of the erythrocyte binding rhoptry protein complex. Trop
Med Int Health 2011, 39:77–82.
70. Fu YX, Li WH: Statistical tests of neutrality of mutations. Genetics 1993,
133:693–709.
71. Hodder AN, Crewther PE, Matthew ML, Reid GE, Moritz RL, Simpson RJ,
Anders RF: The disulfide bond structure of Plasmodium apical membrane
antigen-1. J Biol Chem 1996, 271:29446–29452.
72. Kocken CH, Narum DL, Massougbodji A, Ayivi B, Dubbeld MA, van der Wel
A, Conway DJ, Sanni A, Thomas AW: Molecular characterisation of
Plasmodium reichenowi apical membrane antigen-1 (AMA-1), comparison
with P. falciparum AMA-1, and antibody-mediated inhibition of red cell
invasion. Mol Biochem Parasitol 2000, 109:147–156.
73. Srinivasan P, Beatty WL, Diouf A, Herrera R, Ambroggio X, Moch JK, Tyler JS,
Narum DL, Pierce SK, Boothroyd JC, Haynes JD, Miller LH: Binding of
Plasmodium merozoite proteins RON2 and AMA1 triggers commitment
to invasion. Proc Natl Acad Sci U S A 2011, 108:13275–13280.
74. Mardani A, Keshavarz H, Heidari A, Hajjaran H, Raeisi A, Khorramizadeh MR:
Genetic diversity and natural selection at the domain I of apical
membrane antigen-1 (AMA-1) of Plasmodium falciparum in isolates from
Iran. Exp Parasitol 2012, 130:456–462.
75. Bai T, Becker M, Gupta A, Strike P, Murphy VJ, Anders RF, Batchelor AH:
Structure of AMA1 from Plasmodium falciparum reveals a clustering of
polymorphisms that surround a conserved hydrophobic pocket. Proc
Natl Acad Sci U S A 2005, 102:12736–12741.
76. Conway DJ: Molecular epidemiology of malaria. Clin Microbiol Rev 2007,
20:188–204.
77. Farias MEM, Atkinson CT, LaPointe DA, Jarvi SI: Analysis of the trap gene
provides evidence for the role of elevation and vector abundance in the
genetic diversity of Plasmodium relictum in Hawaii. Malar J 2012,
11:305.
78. Jarvi SI, Farias ME, Atkinson CT: Genetic characterization of Hawaiian
isolates of Plasmodium relictum reveals mixed-genotype infections. Biol
Direct 2008, 3:25.
79. Lamarque M, Besteiro S, Papoin J, Roques M, Vulliez-Le Normand B, Morlon-
Guyot J, Dubremetz J-F, Fauquenoy S, Tomavo S, Faber BW, Kocken CH,
Thomas AW, Boulanger MJ, Bentley GA, Lebrun M: The RON2-AMA1
interaction is a critical step in moving junction-dependent invasion by
Apicomplexan parasites. PLoS Pathog 2011, 7:e1001276.
80. Collins CR, Withers-Martinez C, Hackett F, Blackman MJ: An inhibitory anti-
body blocks interactions between components of the malarial invasion
machinery. PLoS Pathog 2009, 5:e1000273.
81. Richard D, MacRaild CA, Riglar DT, Chan J-A, Foley M, Baum J, Ralph SA,
Norton RS, Cowman AF: Interaction between Plasmodium falciparum
apical membrane antigen 1 and the rhoptry neck protein complex
defines a key step in the erythrocyte invasion process of malaria
parasites. J Biol Chem 2010, 285:14815–14822.
Lauron et al. Malaria Journal 2014, 13:382 Page 13 of 14
http://www.malariajournal.com/content/13/1/382
82. Kato K, Mayer DCG, Singh S, Reid M, Miller LH: Domain III of Plasmodium
falciparum apical membrane antigen 1 binds to the erythrocyte
membrane protein Kx. Proc Natl Acad Sci U S A 2005, 102:5552–5557.
83. Mueller MS, Renard A, Boato F, Vogel D, Naegeli M, Zurbriggen R, Robinson
JA, Pluschke G: Induction of parasite growth-inhibitory antibodies by a
virosomal formulation of a peptidomimetic of loop I from domain III of
Plasmodium falciparum apical membrane antigen 1. Infect Immun 2003,
71:4749–4758.
84. Andreina Pacheco M, Ryan EM, Poe AC, Basco L, Udhayakumar V, Collins
WE, Escalante AA: Evidence for negative selection on the gene encoding
rhoptry –associated protein 1 (RAP-1) in Plasmodium spp. Infect Genet
Evol 2010, 10:655–661.
85. Escalante AA, Cornejo OE, Freeland DE, Poe AC, Durrego E, Collins WE, Lal
AA: A monkey’s tale: The origin of Plasmodium vivax as a human malaria
parasite. Proc Natl Acad Sci U S A 2005, 102:1980–1985.
86. Krief S, Escalante AA, Pacheco MA, Mugisha L, André C, Halbwax M, Fischer
A, Krief JM, Kasenene JM, Crandfield M, Cornejo OE, Chavatte J-M, Lin C,
Letourneur F, Grüner AC, McCutchan TF, Rénia L, Snounou G: On the
diversity of malaria parasites in African Apes and the origin of
Plasmodium falciparum from Bonobos. PLoS Pathog 2010, 6:e1000765.
87. Oliveira DA, Udhayakumar V, Bloland P, Shi YP, Nahlen BL, Oloo AJ, Hawley
WE, Lal AA: Genetic conservation of the Plasmodium falciparum apical
membrane antigen-1 (AMA-1). Mol Biochem Parasitol 1996, 76:333–336.
88. Polley SD, Conway DJ: Strong diversifying selection on domains of the
Plasmodium falciparum apical membrane antigen 1 gene. Genetics 2001,
158:1505–1512.
89. Lal AA, Hughes MA, Oliveira DA, Nelson C, Bloland PB, Oloo AJ, Hawley WE,
Hightower AW, Nahlen BL, Udhayakumar V: Identification of T-cell
determinants in natural immune responses to the Plasmodium
falciparum apical membrane antigen (AMA-1) in an adult population
exposed to malaria. Infect Immun 1996, 64:1054–1059.
90. Şekercioğlu ÇH, Daily GC, Ehrlich PR: Ecosystem consequences of bird
declines. Proc Natl Acad Sci U S A 2004, 101:18042–18047.
91. Tompkins DM, Gleeson DM: Relationship between avian malaria
distribution and an exotic invasive mosquito in New Zealand. J Roy Soc
New Zeal 2006, 36:51–62.
doi:10.1186/1475-2875-13-382
Cite this article as: Lauron et al.:Transcriptome sequencing and analysis
of Plasmodium gallinaceum reveals polymorphisms and selection on the
apical membrane antigen-1. Malaria Journal 2014 13:382.
Submit your next manuscript to BioMed Central
and take full advantage of:
• Convenient online submission
• Thorough peer review
• No space constraints or color figure charges
• Immediate publication on acceptance
• Inclusion in PubMed, CAS, Scopus and Google Scholar
• Research which is freely available for redistribution
Submit your manuscript at
www.biomedcentral.com/submit
Lauron et al. Malaria Journal 2014, 13:382 Page 14 of 14
http://www.malariajournal.com/content/13/1/382
