SciELO - Scientific Electronic Library Online

 
vol.18 número6Evidence of a role for prolactin as regulators of ovarian follicular development in gooseActive anti-acetylcholinesterase component of secondary metabolites produced by the endophytic fungi of Huperzia serrata índice de autoresíndice de materiabúsqueda de artículos
Home Pagelista alfabética de revistas  

Servicios Personalizados

Revista

Articulo

Indicadores

Links relacionados

  • En proceso de indezaciónCitado por Google
  • No hay articulos similaresSimilares en SciELO
  • En proceso de indezaciónSimilares en Google

Compartir


Electronic Journal of Biotechnology

versión On-line ISSN 0717-3458

Electron. J. Biotechnol. vol.18 no.6 Valparaíso nov. 2015

http://dx.doi.org/10.1016/j.ejbt.2015.07.003 

RESEARCH ARTICLE

Characterization of the acetohydroxyacid synthase multigene family in the tetraploide plant Chenopodium quinoa

 

Camilo Mestanzaa, b, Ricardo Riegelc*, Herman Silvad, Santiago C. Vásqueza, e

a Escuela de Graduados, Facultad de Ciencias Agrarias, Universidad Austral de Chile, Valdivia, Chile
b Universidad Técnica Estatal de Quevedo, Quevedo, Ecuador
c Instituto de Producción y Sanidad Vegetal, Facultad de Ciencias Agrarias, Universidad Austral de Chile, Valdivia, Chile
d Departamento de Producción Agrícola, Laboratorio de Genómica Funcional & Bioinformática, Facultad de Ciencias Agronómicas, Universidad de Chile, Santiago, Chile
e Universidad Politécnica Salesiana, Cuenca, Ecuador


ABSTRACT

Background

Currently, the technology called Clearfield® is used in the development of crops resistant to herbicides that inhibit the enzyme acetohydroxy acid synthase (AHAS, EC 2.2.1.6). AHAS is the first enzyme of the biosynthetic pathway that produces the branched-chain of the essential amino acids valine, leucine, and isoleucine. Therefore, multiple copies of the AHAS gene might be of interest for breeding programs targeting herbicide resistance. In this work, the characterization of the AHAS gene was accomplished for the Chenopodium quinoa Regalona-Baer cultivar. Cloning, sequencing, and Southern blotting were conducted to determine the number of gene copies.

Results

The presence of multiple copies of the AHAS gene as has been shown previously in several other species is described. Six copies of the AHAS gene were confirmed with Southern blot analyses. CqHAS1 and CqAHAS2 variants showed the highest homology with AHAS mRNA sequences found in the NR Database. A third copy, CqAHAS3, shared similar fragments with both CqAHAS1 and CqAHAS2, suggesting duplication through homeologous chromosomes pairing.

Conclusions

The presence of multiple copies of the gene AHAS shows that gene duplication is a common feature in polyploid species during evolution. In addition, to our knowledge, this is the first report of the interaction of sub-genomes in quinoa.

Keywords: Acetohydroxyacid synthase; Acetolactate synthase; Chenopodium quinoa; Homeologous pairing; Gene duplication


 

1. Introduction

Quinoa (Chenopodium quinoa Willd) is an allotetraploid plant (2n = 4 × = 36) [1], that is native to the Andean region of South America, and has been cultivated for thousands of years [2]. Recent international attention given to quinoa is centered on its unusually high nutritional value [3]. Compared with cereals, quinoa's protein content is higher (14 to 16%) and provides an ideal balance of essential amino acids for human consumption [4]. Despite these properties, there has been a lack of research and support for quinoa to the point that it is considered a neglected crop [5]. The quinoa crop in northern Chile is cultivated primarily by indigenous Aymara in the Chilean Altiplano; nevertheless, the cultivation of quinoa extends to the south-central zone of Chile in a fragmented pattern. Chilean quinoa is characterized by a broad range of morphological diversity that likely resulted from artificial selection, natural selection, and genetic drift as landraces were introduced to south-central Chile via the trade and migration of indigenous peoples [6].

The productive potential of the Regalona-Baer cultivar (that belongs to a group of coastal ecotypes) in Chile is very high and comparable to the yield of rice (6.2 t ha- 1) [7]. This variety of quinoa produces yields of 3 t ha- 1 and can even produce over 8 t ha- 1 in small plots [8]. The world production of quinoa has increased over the last ten years mostly due to a rise in cultivated land and not by improvements in production per hectare [7]. The huge gap between the real and the potential yield of this crop generates a spiral of negative effects that restrict the expansion of this valuable crop. Probably, the most important constraint on the industrialized production of crops is weed control [9]. This is also true for quinoa production where weed control has a major impact and is one of the main factors that has limited quinoa expansion [10]. Weeds may have a major impact on seed yield. For instance, it has been shown that the yield may be tripled if grass weeds are controlled [11]. After quick emergence, further development is slow, and quinoa may be over grown by weeds [12]. So far, there are no registered herbicides for quinoa, and in addition, C. quinoa is a crop species for which problem weeds include closely related members of the family Amaranthaceae [13], making classical herbicide selection difficult. The development of herbicide-resistant crops has resulted in significant changes in agronomic practices, one of which is the adoption of effective, simple, low-risk, crop-production systems with less dependency on tillage and lower energy requirements [14].

In recent years, different companies have conducted extensive research and technological development through public-private partnerships in order to develop crops resistant to herbicides. Commercially, one of the products developed using this approach is called Clearfield®. By 2004, five imidazolinone-tolerant crops, in combination with four imidazolinone herbicides, have been commercialized in different regions of the world and make up what is called the Clearfield® production system [15]. This technology is based on the resistance to herbicides that inhibit the enzyme acetohydroxy acid synthase (AHAS, EC 2.2.1.6). AHAS, is the first enzyme necessary for the biosynthetic pathway that produces the branched-chain essential amino acids valine, leucine, and isoleucine [16] and [17]. AHAS is also the common target site of five herbicide chemical groups: sulfonylurea (SU), imidazolinone (IMI), triazolopyrimidine (TP), pyrimidinyl-thiobenzoates (PTB) and sulfonyl-aminocarbonyl-triazolinone (SCT). Inhibition of AHAS starves plants of valine, leucine, and isoleucine, which leads to plant death [18], [19] and [20].

Depending on the ploidy level, individuals may have a variable number of AHAS gene copies and these copies can be more complex than those found in diploid organisms [21] and [22]. Multiple AHAS genes have been reported in many plant species belonging to different genera and having different ploidy [23] and [24]. Regardless of the high nutritional and economic potential of C. quinoa, there are few studies related to increasing the cultivation and yield of this crop. The first step to increase production would be to achieve efficient weed control. In light of this, the aim of this study is to characterize the gene family encoding the AHAS enzyme in the quinoa Regalona-Baer cultivar to provide the basic information needed for a herbicide resistant breeding program.

2. Materials and methods

2.1. Plant material and DNA extraction

For the extraction of genomic DNA, a unique assortment of quinoa cultivars registered in Chile (Regalona-Baer) was used. Seeds were sown in 300 ml pots with a mixture of soil/sand and were grown using the following greenhouse conditions: 20/12°C day/night (± 4°C) with a light regime of 14/10 h day/night and 900 µmol m- 2 s- 1 light intensity. Leaf tissue was harvested when plants had four true leaves and DNA extraction was performed using the CTAB modified protocol [25]. DNA was quantified by absorbance at 260/280 nm [26]. Extracts were diluted in distilled water (20 ng µl- 1) and stored at -20°C for future analysis.

2.2. AHAS gene discovery

To select and design primers, existing information from the NCBI GenBank database on the closely related genus Amaranthus, Bassia and Salsola was used. For sequence alignment, Amaranthus hypochondriacus, Amaranthus tuberculatus, Amaranthus retroflexus, Amaranthus powellii, Bassia scoparia and Salsola tragus were used. Conserved regions were identified and primers were designed using the Primer3 software [27] (Table 1). The PCR mix consisted of 2 µl of total DNA (20 ng µl- 1), 2 µl of each primer (10 ?M), 4 µl of MgCl2 (25 mM), 2 µl of each dNTP (2.5 mM), 2 units of Platinum Taq DNA polymerase and 5 µl of 10 × buffer supplied with the enzyme, in a final volume of 50 µl. The amplification protocol consisted of 3 min incubation at 94°C, 34 cycles of 35 s at 94°C, 45 s at x°C, and 105 s at 72°C, and a final extension of 7 min at 72°C. x°C represents the annealing temperature for each pair of primers used (Table 1 and Table 2).

Table 1. Primers used for the amplification of the AHAS gene sequence.

Primer Nucleotide sequence (5'-3') Genus
F10
CCTAAACCTAAACCTCCTTC
Amaranthus
F1
TTTTGTTTCCCGATTTAGCCC
Amaranthus
F3
ATTCCTCCGCAATACGCCATT
Amaranthus
R4
AATCAAAACAGGTCCAGGTC
Amaranthus
R1
CCTACAAAAAGCTTCTCCTCTATAAG
Amaranthus
RUTH-F-1C
CKGGCCGTGTKGGTGTTTG
Salsola
RUTH-R-3B
AACTTGTTCTTCCATCACCTTCG
Salsola
B1-F
ATGGCGTCTACTTGTGCAAATCC
Bassia
CHALSF1
GCGTCTACTTGTKCAAAYC
Chenopodium
CHALSF4
GACCTGGACCTGTTTTGATT
Chenopodium
CHALS R1
CAAAGTAACAAGCAACATRAMAAC
Chenopodium
ALS1FB*
ATCACCCCTTCTCTTCTTCAA
Chenopodium
ALS1RD*
CAACAACAAACTAACCTAAAGCA
Chenopodium
ALSGR1*
CATCAAACCTAACCCCGAAA
Chenopodium
ALS2RD*
AGTAGTAGCAAGCAGCATGTG
Chenopodium
ALSGF2*
TTTCGGGGTTAGGTTTGATG
Chenopodium
ALS2F*
ACAAAAATAAACCCTACTCCGTA
Chenopodium
*Copy-specific amplification.

 

Table 2. Results of the combinations of primers used for amplification of the AHAS gene sequence.

Combination Amplification
F10-R4
Unspecific
F1-R4 (Fragment B)
Positive at 58°C
F1-CHALSR1
Unspecific
CHALSF1-R4 (Fragment A)
Positive to 57°C
F3-CHALSR1
Unspecific
CHALSF4-R1
Unspecific
CHALSF4-CHALSR1
Unspecific
B1-F-CHALSR1
Unspecific
B1-F-R4
Unspecific
F1-RUTH-R-3B (Fragment E)
Positive at 58°C
CHALSF4-RUTH-R-3B (Fragment D)
Positive at 57°C
RUTH-F-1C-RUTH-R-3B (Fragment C)
Positive at 60°C
*ALS1FB-ALSGR1
Positive at 58°C
*ALS2F-ALSGR1
Positive at 58°C
*ALSGF2-ALS1RD
Positive at 58°C
*ALSGF2-ALS2RD
Positive at 58°C
* Copy-specific amplification.

 

Amplified products were analyzed by gel electrophoresis using a 1.2% agarose gel prepared with 1 × TAE buffer and ethidium bromide 0.5 µg ml- 1. The gel was run for 30 min at 100 V. DNA bands were visualized under UV light, compared with a molecular weight marker, and photographed. Fragments of interest were then sequenced by Macrogen, Korea. Confirmation of identity and consensus sequence identification was performed with nucleotide Blast and BlastX of NCBI. The transit peptide of AHAS was identified using ChloroP and TargetP 1.1 [28].

To determine the number of gene copies, the kit pGEMT Easy Vector kit (Promega) was used for cloning the fragment B (499 bp) (Table 2) following all manufacturer's instructions. This fragment was selected because it showed the clearest chromatograms and contains three of the five conserved protein domains. Thirty colonies positive for each transformation were selected and amplified using primers SP6 and T7. Following this, sequencing was conducted for both strand directions; this stage was made by Macrogen, Korea. To infer the phylogenetic relationship of the sequenced genes, phylogenetic trees were constructed using the program MEGA with the statistical methods of maximum parsimony and Bootstrapping using 1000 replicates [29].

Furthermore, a Southern blot analysis was done using the protocol of Sambrook [30]. The FastPCR program was used to search enzymes that do not cut the sequence of the probe used (BspTI and SspI), and as control, enzymes that cut one copy (Copy 2) (Mph1103I), and enzymes that cut all copies (EcoRI). For hybridization and detection, the DecaLabel Biotin Labeling Kit™ DNA Detection Kit and Chromogenic Biotin were used following the instructions of the manufacturer. The analysis of bands was performed with the UN-SCAN-IT gel 6.1 software; and DNA bands were visualized under UV light, compared with a molecular weight marker, and photographed.

The new sequences registered for quinoa were compared with quinoa AHAS mRNA obtained from aerial and root tissue of seedlings with 36 days of growth, that have been used for analyzing the transcriptome of quinoa (#2238, KM233690 and #2237, KM233691; Ecotype R49; Morales, Zurita and Silva, unpublished results), using the program MEGA with the statistical methods of maximum parsimony and Bootstrapping using 1000 replicates [29].

3. Results

3.1. Isolation and identification of AHAS in C. quinoa

Five primer combinations were used for direct sequencing since they contain the region spanning the transit peptide and the domains C, A, D, B and E (Table 2). The best amplifications were obtained with fragments B and D. The PCR fragments in total contained 1992 bp, not including the start region and the stop codon.

The confirmation of identity of the new fragment of quinoa using the BLASTN [31] from NCBI, exhibited E-values of 0,0; and an 87% of identity with B. scoparia, 86% with S. tragus, and 84% with A. powellii, A. retroflexus and A. tuberculatus. Analyses with ChloroP and TargetP 1.1 show that the transit peptide has an estimated size of 50 amino acids, with a score of 0.989 for prediction of localization in the chloroplast.

The complete AHAS gene sequence has 2001 bp and does not contain introns; with a total of 667 amino acids including the start and stop codons. The phylogeny made using maximum parsimony grouped the AHAS sequence of C. quinoa within the subfamily Chenopodiaceae, family Amaranthaceae (Fig. 1).

Fig. 1. Hypothetical phylogenetic relationships of plants inferred by maximum parsimony. AHAS sequences including the new sequence of Chenopodium quinoa (CqAHAS1) inferred by maximum parsimony. The oval shows the Chenopodiaceae subfamily members. The numbers at the nodes represent bootstrap values (1000 replicates). Amaranthus powellii, Amaranthus retroflexus, Amaranthus hypochondriacus, Amaranthus tuberculatus, Salsola tragus, Bassia scoparia, Conyza canadensis, Anthemis cotula, Arabidopsis thaliana, Brassica napus, Zea mays.

 

3.2. Sequence variation in the AHAS gene of C. quinoa

Analysis of the chromatograms obtained from sequencing performed by Macrogen revealed double peaks in the same position in the chromatogram, suggesting the presence of more than one copy of the AHAS gene (data not shown). The presence of the AHAS gene family in polyploid quinoa was confirmed by cloning and sequencing fragment B which indicated the presence of six copies: CqAHAS1, CqAHAS2, CqAHAS3, CqAHAS4, CqAHAS5 and CqAHAS6 (Fig. 2). These six copies were validated using a Southern blot (Fig. 3).

Fig. 2. Hypothetical phylogenetic relationships among AHAS gene copies in Chenopodium quinoa inferred by maximum parsimony. The numbers at the nodes represent bootstrap values (1000 replicates). CqAHAS1, CqAHAS2, CqAHAS3, CqAHAS4, CqAHAS5, CqAHAS6, 2238, 2237 and Salsola tragus.

Fig. 3. Southern blot analyses using endonucleases Mph1103I, EcoRI, SspI and BspTI, six copies of the AHAS gene of Chenopodium quinoa Regalona-Baer variety were detected. M represents a molecular marker in kilo bases. The digitalized membrane indicates the number of detected bands using UN-SCAN-IT gel 6.1 software.

 

The copies of the AHAS gene were compared with mRNA sequences #2238, KM233690 and #2237, KM233691. The analysis of the hypothetical phylogenetic relationships AHAS gene copies in C. quinoa inferred by maximum parsimony formed two well-delineated groups: copy 1 and copy 4 had 99.7 and 99.5% identity with sequence #2238, respectively. While, copies 2, 5, and 6 showed 99.7, 99.5, and 99.3% identity with sequence #2237, respectively (Fig. 2). The CqAHAS3 copy is mainly separated from the two groups and most likely lies between the two groups.

In addition, we isolated the complete sequence of the most representative (CqAHAS1, KM253767 and CqAHAS2, KM253768) copies of the Regalona-Baer variety. Based on these sequences, we designed specific primers for each gene copy (Table 1). Expected PCR products were obtained and direct sequencing was possible and feasible. This procedure will allow us to conduct, in the future, TILLING and ECO-TILLING analyses for the identification of putative point mutations and/or early selection based on an allele-specific analysis of natural and mutant populations of quinoa [32].

Interestingly, when looking at the results of the CqAHAS3 copy in the grouping, we analyzed the nucleotide sequence, finding an unexpected result: the initial part of the segment (133 bp) was similar to CqAHAS1, while the rest of the segment (348 bp) was more similar to CqAHAS2 (Fig. 4).

Fig. 4. Partial nucleotide sequences of CqAHAS1, CqAHAS2 and CqAHAS3. Polymorphic sites are colored and similar colors show homology among sequences.

4. Discussion

The 2001 bp sequencing encoding the AHAS gene in C. quinoa is similar to that of Bassia while it differs from Amaranthus. The AHAS gene in Amaranthus is 2010 bp long, and the differences in the gene size could be due to polymorphic indels in the C- and N-terminal regions of the protein [17]. Nevertheless, these terminal regions are involved in intracellular traffic rather than in the catalytic activity of the enzyme [17].

Despite these mentioned differences, the 50-aa transit peptide and localization cellular of C. quinoa is similar to that of A. tuberculatus [33], but differs from the three AHAS copies of Brassica napus [34] and Arabidopsis thaliana [35]. Furthermore, the AHAS gene sequence of C. quinoa, with a total of 667 amino acids without the presence of introns, is similar to that of many monocots and dicots [36], [37], [38], [39], [40], [41], [42], [43], [44], [45], [46], [47], [48], [49], [50], [51] and [52]. Exceptional cases are Lindernia spp. [53], Schoenoplectus juncoides [54], and Schoenoplectus mucronatus [55] whose sequences possess introns and therefore have alternative splicing.

The six variants of the AHAS gene found in this study were unexpected for C. quinoa as diploid species of the Amaranthaceae family, are thought to have only one copy of this gene [43]. Only the tetraploid species Salsosa tragus exhibits two variants [39]. The maximum parsimony phylogenies integrating sequences obtained by cloning (481 bp) and mRNA sequences retrieved from databases, suggest that only CqAHAS1 and CqAHAS2 are functional copies. It is possible that each of these two copies derived from the different diploid parental ancestor and underwent gene duplication after hybridization and polyploidization. Regarding the CqAHAS3 copy, the initial part of the segment (133 bp) was similar to CqAHAS1 however the rest of the segment (348 bp) was more similar to CqAHAS2 (Fig. 4). This suggests homeologous chromosome pairing assuming that CqAHAS1 and CqAHAS2 come from different diploid ancestors. In fact, Pecinka et al. [56] point out that in polyploids, an increase in the frequency of meiotic recombination (due to pairing and sorting of not only more than two homeologous chromosomes, but also of the homeologous chromosomes during meiosis) could boost the amount of genetic diversity, thereby enhancing the adaptation potential of the organism in challenging natural environments.

The third copy and other variants (4, 5 and 6) of the AHAS gene found in C. quinoa seem to be silent or might be expressed in a specific tissue. Further gene expression studies are needed to corroborate this. The presence of multiple copies of AHAS in the active, silenced, sub-functionalized or neo-functionalized form is well documented in databases particularly for allotetraploid species [21], [39] and [57]. Based on this, polyploids typically show a redistribution and/or increase in the number of loci [58] as a product of gene duplications [59]. The complete sequencing of several eukaryotic genomes shows the importance of duplicated genes [60] and [61]. In particular, plant genomes contain a high proportion of duplicated genes that are also often highly redundant. In several plants, there are more than one hundred documented paralogous genes [62] and [63].

The results of this came as part of the complexity of the quinoa tetraploid genome, reflecting, as in other studies, the allotetraploidy in quinoa [64], [65], [66] and [67], with functional alleles having been retained at some duplicated loci and at least, some association occurring between homeologous chromosomes, product of erratic multivalent formation at meiosis [64]. Different situation happens with other genes, such as the Salt Overly Sensitive 1 (SOS1), where Maughan et al. [68] suggest possible conservation of synteny across the C. quinoa sub-genomes across the homeologous SOS1.

Multiple copies of the AHAS gene are probably not necessary for the growth and development of the species [69]. However, in mutation breeding programs, having multiple copies of AHAS genes could have a positive effect on herbicide resistance [22], [70] and [71]. Multiple copies of a gene, product of its polyploidy, bring benefits naturally, without generating gene stacking with transgenic techniques [72].

In this context, higher levels of resistance, due to an increased number of copies of the gene and resistance additive effect [22] and [71], are desirable to ensure minimal plant injury at herbicide addition rates that are required for adequate weed control [73]. Higher levels of resistance to AHAS inhibiting herbicides have been observed in polyploid species when multiple resistance alleles are present [70]. It has been described that alleles conferring imidazolinone tolerance in polyploid species has an additive genetic effect [15]. In the allohexaploid bread wheat, mutations in two copies of the AHAS gene allow for an increase in the level of herbicide resistance compared with genotypes having only one mutated gene [22]. This work shows the first molecular characterization of the AHAS genes in quinoa as well as the interaction between its homeologous genomes.

Financial support

This thesis project is associated with CONICYT Program No. 7813110011 and CONICYT, Scientific Information Program/Fund for Scientific Journals Publishing, Year 2014, ID FP140010.

Data deposition: The sequences reported in this work for quinoa have been deposited in the Gen Bank database (accession nos. KM233692, KM233693, KM233694, KM233695, KM233696, KM233697, KM253767 and KM253768). The other sequences used are: Amaranthus powellii (AF363370.1), Amaranthus retroflexus (AF363369.1), Amaranthus hypochondriacus (EU024568.1), Amaranthus tuberculatus (EF157818.1), Salsola tragus (GU271180.1), Bassia scoparia (EU517465.1), Conyza canadensis (HM067014.1), Anthemis cotula (JF327752.1), Arabidopsis thaliana (NM_114714.2), Brassica napus (GU192448.1), and Zea mays (NM_001158289.1).

Acknowledgments

The authors thank SENESCYT-Ecuador and the Escuela de Graduados of the Facultad de Ciencias Agrarias of the Universidad Austral de Chile. Jaime Figueroa and Judith Carrasco are thanked for their assistance in the laboratory. Thanks are given to Milton Gallardo Narcisi for his valuable assistance in the preparation and review of this manuscript.

References

1. Giusti L. El género Chenopodium en Argentina: I. Números de cromosomas. Darwiniana 1970;16:98-105.         [ Links ]

2. D'Altroy TN, Hastorf CA. The distribution and contents of Inca state storehouses in the Xauxa region of Peru. Am Antiq 1984;49:334-49.         [ Links ]

3. Jacobsen SE. The worldwide potential for quinoa (Chenopodium quinoa Willd.). Food Rev Int 2003;19:167-77. http://dx.doi.org/10.1081/FRI-120018883.         [ Links ]

4. Kozioł MJ. Chemical composition and nutritional evaluation of quinoa (Chenopodium quinoa Willd.). J Food Compos Anal 1992;5:35-68. http://dx.doi.org/10.1016/0889-1575(92)90006-6.         [ Links ]

5. Rojas W, Valdivia R, Padulosi S, Pinto M, Soto JL, Alcócer E, et al. From neglect to limelight: Issues, methods and approaches in enhancing sustainable conservation and use of Andean grains in Bolivia and Peru. J Agric Rural Dev Trop Subtrop 2009;92:87-117.         [ Links ]

6. Fuentes FF, Martinez E, Hinrichsen PV, Jellen EN, Maughan PJ. Assessment of genetic diversity patterns in Chilean quinoa (Chenopodium quinoa Willd.) germplasm using multiplex fluorescent microsatellite markers. Conserv Genet 2008;10:369-77. http://dx.doi.org/10.1007/s10592-008-9604-3.         [ Links ]

7. FAO. FAO-FAOSTAT 2014. http://faostat.fao.org/site/567/DesktopDefault.aspx?PageID=567#ancor. [cited July 23, 2014. Available from Internet at:].         [ Links ]

8. Von Baer I, Bazile D, Martinez E. Cuarenta años de mejoramiento de quínoa (Chenopodium quinoa Willd.) en la Araucanía: Origen de "La Regalona-B.". Rev Geogr Valparaíso 2009;42:34-44.         [ Links ]

9. Chauhan BS. Weed ecology and weed management strategies for dry-seeded rice in Asia. Weed Technol 2012;26:1-13. http://dx.doi.org/10.1614/WT-D-11-00105.1.         [ Links ]

10. Jacobsen S-E, Christiansen JL, Rasmussen J.Weed harrowing and inter-row hoeing in organic grown quinoa (Chenopodium quinoa Willd.). Outlook Agric 2010;39:223-7. http://dx.doi.org/10.5367/oa.2010.0001.         [ Links ]

11. Johnson DL, Ward SM. Quinoa. In: Janick, Simon JE, editors. New York: New Crop; 1993. p. 219-21.         [ Links ]

12. Ruiz KB, Biondi S, Oses R, Acuña-Rodríguez IS, Antognoni F, Martinez-Mosqueira E, et al. Quinoa biodiversity and sustainability for food security under climate change. A review. Agron Sustain Dev 2013;34:349-59. http://dx.doi.org/10.1007/s13593-013-0195-0.         [ Links ]

13. Coile NC, Artaud CR. Chenopodium ambrosioides L., (Chenopodiaceae). Bot Circ 1998: 1-6.         [ Links ]

14. Vencill WK, Nichols RL, Webster TM, Soteres JK, Mallory-Smith C, Burgos NR, et al. Herbicide resistance: Toward an understanding of resistance development and the impact of herbicide-resistant crops. Weed Sci 2012;60:2-30. http://dx.doi.org/10.1614/WS-D-11-00206.1.         [ Links ]

15. Tan S, Evans RR, Dahmer ML, Singh BK, Shaner DL. Imidazolinone-tolerant crops: History, current status and future. Pest Manag Sci 2005;61:246-57. http://dx.doi.org/10.1002/ps.993.         [ Links ]

16. Duggleby RG, Mccourt JA, Guddat LW. Structure and mechanism of inhibition of plant acetohydroxyacid synthase. Plant Physiol Biochem 2008;46:309-24. http://dx.doi.org/10.1016/j.plaphy.2007.12.004.         [ Links ]

17. Duggleby RG, Pang SS. Acetohydroxyacid synthase. J Biochem Mol Biol 2000;33: 1-36.         [ Links ]

18. Powles SB, Yu Q. Evolution in action: Plants resistant to herbicides. Annu Rev Plant Biol 2010;61:317-47. http://dx.doi.org/10.1146/annurev-arplant-042809-112119.         [ Links ]

19. Singh BK, Stidham MA, Shaner DL. Assay of acetohydroxyacid synthase. Anal Biochem 1988;171:173-9. http://dx.doi.org/10.1016/0003-2697(88)90139-X.         [ Links ]

20. Shaner DL, Anderson PC, Stidham MA, Company AC, Box P. Imidazolinones: Potent inhibitors of acetohydroxyacid synthase. Plant Physiol 1984;76:545-6.         [ Links ]

21. Grula JW, Hudspeth RL, Hobbs SL, Anderson DM. Organization, inheritance and expression of acetohydroxyacid synthase genes in the cotton allotetraploid Gossypium hirsutum. Plant Mol Biol 1995;28:837-46.         [ Links ]

22. Pozniak CJ, Hucl PJ. Genetic analysis of imidazolinone resistance inmutation-derived lines of common wheat. Crop Sci 2004;44:23-30.         [ Links ]

23. Panozzo S, Scarabel L, Tranel PJ, Sattin M. Target-site resistance to ALS inhibitors in the polyploid species Echinochloa crus-galli. Pestic Biochem Physiol 2013;105: 93-101. http://dx.doi.org/10.1016/j.pestbp.2012.12.003.         [ Links ]

24. Riar DS, Norsworthy JK, Bond J, Bararpour MT, Wilson MJ, Scott RC. Resistance of Echinochloa crus-galli populations to acetolactate synthase-inhibiting herbicides. Int, J Agron 2012;2012:1-8. http://dx.doi.org/10.1155/2012/893953.         [ Links ]

25. Arismendi N, Andrade N, Riegel R, Carrillo R. Presence of a phytoplasma associated with witches' broom disease in Ugni molinae Turcz. and Gaultheria phillyreifolia (Pers.) Sleumer determined by DAPI, PCR, and DNA sequencing. Chil J Agric Res 2010;70:26-33.         [ Links ]

26. Shokere L, Holden MJ, Ronald Jenkins G. Comparison of fluorometric and spectrophotometric DNA quantification for real-time quantitative PCR of degraded DNA. Food Control 2009;20:391-401. http://dx.doi.org/10.1016/j.foodcont.2008.07.009.         [ Links ]

27. Rozen S, Skaletsky H. Primer3 on the WWW for general users and for biologist programmers. Methods Mol Biol 2000;132:365-86.         [ Links ]

28. 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;8:978-84. http://dx.doi.org/10.1110/ps.8.5.978.         [ Links ]

29. Bhattacharya D. Analysis of the distribution of bootstrap tree lengths using the maximum parsimony method. Mol Phylogenet Evol 1996;6:339-50. http://dx.doi.org/10.1006/mpev.1996.0084.         [ Links ]

30. Sambrook J, Fritsch E, Maniatis T. Molecular cloning. A laboratory manual. Vol. 12nd ed. ; 1989[New York].         [ Links ]

31. Zhang Z, Schwartz S, Wagner L, Miller W. A greedy algorithm for aligning DNA sequences. J Comput Biol 2000;7:203-14.         [ Links ]

32. Till BJ, Reynolds SH, Greene EA, Codomo CA, Enns LC, Johnson JE, et al. Large-scale discovery of induced point mutations with high-throughput TILLING. Genome Res 2003;13:524-30. http://dx.doi.org/10.1101/gr.977903.         [ Links ]

33. Patzoldt WL, Tranel PJ. Multiple ALS mutations confer herbicide resistance in waterhemp (Amaranthus tuberculatus). Weed Sci 2007;55:421-8. http://dx.doi.org/10.1614/WS-06-213.1.         [ Links ]

34. Rutledge RG, Hattori J, Miki BL. Molecular characterization and genetic origin of the Brassica napus acetohydroxyacid synthase multigene family. Mol Gen Genet 1991; 229:31-40.         [ Links ]

35. Sathasivan K, Haughn GW, Murai N. Nucleotide sequence of a mutant acetolactate synthase gene from an imidazolinone-resistant Arabidopsis thaliana var. Columbia. Nucleic Acids Res 1990;18:2188.         [ Links ]

36. Tranel PJ,Wright TR. Review resistance of weeds to ALS-inhibiting herbicides:What have we learned ? Weed Sci 2002;50:700-12.         [ Links ]

37. Shivrain VK, Burgos NR, Sales M, Kuk YI. Polymorphisms in the ALS gene of weedy rice (Oryza sativa L.) accessions with differential tolerance to imazethapyr. Crop Prot 2010;29:336-41. http://dx.doi.org/10.1016/j.cropro.2009.10.002.         [ Links ]

38. Intanon S, Perez-Jones A, Hulting AG, Mallory-Smith C. Multiple pro 197 ALS substitutions endow resistance to ALS inhibitors within and among mayweed chamomile populations. Weed Sci 2011;59:431-7. http://dx.doi.org/10.1614/WS-D-10-00146.1.         [ Links ]

39. Warwick SI, Sauder C, Beckie HJ. Acetolactate synthase (ALS) target-site mutations in ALS inhibitor-resistant russian thistle (Salsola tragus). Weed Sci 2010;58: 244-51. http://dx.doi.org/10.1614/WS-D-09-00083.1.         [ Links ]

40. Délye C, Pernin F, Scarabel L. Evolution and diversity of the mechanisms endowing resistance to herbicides inhibiting acetolactate-synthase (ALS) in corn poppy (Papaver rhoeas L.). Plant Sci 2011;180:333-42. http://dx.doi.org/10.1016/j.plantsci.2010.10.007.         [ Links ]

41. Laplante J, Rajcan I, Tardif FJ. Multiple allelic forms of acetohydroxyacid synthase are responsible for herbicide resistance in Setaria viridis. Theor Appl Genet 2009;119: 577-85. http://dx.doi.org/10.1007/s00122-009-1067-5.         [ Links ]

42. Reith M, Munholland J. Two amino-acid biosynthetic genes are encoded on the plastid genome of the red alga Porphyra umbilicalis. Curr Genet 1993;23:59-65. http://dx.doi.org/10.1007/BF00336751.         [ Links ]

43. Warwick SI, Xu R, Sauder C, Beckie HJ. Acetolactate synthase target-site mutations and single nucleotide polymorphism genotyping in ALS-Resistant Kochia (Kochia scoparia). Weed Sci 2008;56:797-806. http://dx.doi.org/10.1614/WS-08-045.1.         [ Links ]

44. Wright TR, Penner D. Cell selection and inheritance of imidazolinone resistance in sugarbeet (Beta vulgaris). Theor Appl Genet 1998;96:612-20.         [ Links ]

45. Mcnaughton KE, Letarte J, Lee EA, Tardif FJ. Mutations in ALS confer herbicide resistance in redroot pigweed (Amaranthus retroflexus) and Powell amaranth (Amaranthus powellii). Weed Sci 2005;53:17-22.         [ Links ]

46. Sibony M, Michel A, Haas HU, Rubin B, Hurle K. Sulfometuron-resistant Amaranthus retroflexus: Cross-resistance and molecular basis for resistance to acetolactate synthase-inhibiting herbicides. Weed Res 2001;41:509-22.         [ Links ]

47. Scarabel L, Varotto S, Sattin M. A European biotype of Amaranthus retroflexus cross-resistant to ALS inhibitors and response to alternative herbicides. Weed Res 2007;47:527-33. http://dx.doi.org/10.1111/j.1365-3180.2007.00600.x.         [ Links ]

48. Maertens KD, Sprague CL, Tranel PJ, Hines RA. Amaranthus hybridus populations resistant to triazine and acetolactate synthase-inhibiting herbicides. Weed Res 2004;44:21-6.         [ Links ]

49. Trucco F, Hager AG, Tranel PJ. Acetolactate synthase mutation conferring imidazolinone-specific herbicide resistance in Amaranthus hybridus. J Plant Physiol 2006;163:475-9. http://dx.doi.org/10.1016/j.jplph.2005.06.015.         [ Links ]

50. Sibony M, Rubin B. Molecular basis for multiple resistance to acetolactate synthaseinhibiting herbicides and atrazine in Amaranthus blitoides (prostrate pigweed). Planta 2003;216:1022-7. http://dx.doi.org/10.1007/s00425-002-0955-6.         [ Links ]

51. Ferguson GM, Hamill AS, Tardif FJ. ALS inhibitor resistance in populations of Powell amaranth and redroot pigweed. Weed Sci 2001;49:448-53.         [ Links ]

52. Diebold RS,McNaughton KE, Lee E, Tardif FJ. Multiple resistance to imazethapyr and atrazine in Powell amaranth (Amaranthus powellii). Weed Sci 2003;51:312-8. http://dx.doi.org/10.1614/0043-1745(2003)051[0312:MRTIAA]2.0.CO;2.         [ Links ]

53. Uchino A, Watanabe H. Mutations in the acetolactate synthase genes of sulfonylurea-resistant biotypes of Lindernia spp. Weed Biol Manag 2002;2:104-9.         [ Links ]

54. Uchino A, Ogata S, Kohara H, Yoshida S, Yoshioka T, Watanabe H. Molecular basis of diverse responses to acetolactate synthase-inhibiting herbicides in sulfonylurearesistant biotypes of Schoenoplectus juncoides. Weed Biol Manag 2007;7:89-96. http://dx.doi.org/10.1111/j.1445-6664.2007.00240.x.         [ Links ]

55. Scarabel L, Locascio A, Furini A, Sattin M, Varotto S. Characterisation of ALS genes in the polyploid species Schoenoplectus mucronatus and implications for resistance management. Pest Manag Sci 2010;66:337-44. http://dx.doi.org/10.1002/ps.1883.         [ Links ]

56. Pecinka A, Fang W, Rehmsmeier M, Levy A, Mittelsten Scheid O. Polyploidization increases meiotic recombination frequency in Arabidopsis. BMC Biol 2011;9:24. http://dx.doi.org/10.1186/1741-7007-9-24.         [ Links ]

57. Ouellet T, Rutledge RG, Miki BL. Members of the acetohydroxyacid synthase multigene family of Brassica napus have divergent patterns of expression. Plant J 1992;2:321-30.         [ Links ]

58. Weiss-Schneeweiss H, Emadzade K, Jang T-S, Schneeweiss GM. Evolutionary consequences, constraints and potential of polyploidy in plants. Cytogenet Genome Res 2013;140. http://dx.doi.org/10.1159/000351727.Evolutionary.         [ Links ]

59. Ohno S, Wolf U, Atkin NB. Evolution from fish to mammals by gene duplication. Hereditas 1968;59:169-87.         [ Links ]

60. Wapinski I, Pfeffer A, Friedman N, Regev A. Natural history and evolutionary principles of gene duplication in fungi. Nature 2007;449:54-61. http://dx.doi.org/10.1038/nature06107.         [ Links ]

61. Tekaia F, Dujon B. Pervasiveness of gene conservation and persistence of duplicates in cellular genomes. J Mol Evol 1999;49:591-600.         [ Links ]

62. The Arabidopsis genome initiative. Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 2000;408:796-815. http://dx.doi.org/10.1038/35048692.         [ Links ]

63. Yu J, Hu S,Wang J, Dai L, Zhou Y, Zhang X, et al. A draft sequence of the rice genome (Oryza sativa L. ssp. indica). Science 2002;296:79-92. http://dx.doi.org/10.1126/science.1068037.         [ Links ]

64. Ward SM. Allotetraploid segregation for single-gene morphological characters in quinoa (Chenopodium quinoa Willd.). Euphytica 2000;116:11-6.         [ Links ]

65. Maughan PJ, Bonifacio A, Jellen EN, Stevens MR, Coleman CE, Ricks M, et al. A genetic linkage map of quinoa (Chenopodium quinoa) based on AFLP, RAPD, and SSR markers. Theor Appl Genet 2004;109:1188-95. http://dx.doi.org/10.1007/s00122-004-1730-9.         [ Links ]

66. Brown DC, Cepeda-Cornejo V, Maughan PJ, Jellen EN. Characterization of the granule-bound starch synthase I gene in Chenopodium. Plant Genome 2015;8. http://dx.doi.org/10.3835/plantgenome2014.09.0051.         [ Links ]

67. Kolano B, Gardunia BW, Michalska M, Bonifacio A, Fairbanks D, Maughan PJ, et al. Chromosomal localization of two novel repetitive sequences isolated from the Chenopodium quinoa Willd genome. Genome 2011;717:710-7. http://dx.doi.org/10.1139/G11-035.         [ Links ]

68. Maughan PJ, Turner TB, Coleman CE, Elzinga DB, Jellen EN, Morales Ja, et al. Characterization of Salt Overly Sensitive 1 (SOS1) gene homoeologs in quinoa (Chenopodium quinoa Willd.). Genome 2009;52:647-57. http://dx.doi.org/10.1139/G09-041.         [ Links ]

69. Keeler SJ, Sanders P, Smith JK, Mazur BJ. Regulation of tobacco acetolactate synthase gene expression. Plant Physiol 1993;102:1009-18.         [ Links ]

70. Swanson EB, Herrgesell MJ, Arnoldo M, Sippell DW,Wong RS.Microsporemutagenesis and selection: Canola plants with field tolerance to the imidazolinones. Theor Appl Genet 1989;78:525-30. http://dx.doi.org/10.1007/BF00290837.         [ Links ]

71. Pozniak CJ, Birk IT, Donoughue LSO, Me C, Hucl PJ, Singh BK. Physiological and molecular characterization of mutation-derived imidazolinone resistance in spring wheat. Crop Sci 2004;44:1434-43.         [ Links ]

72. Halpin C. Gene stacking in transgenic plants - The challenge for 21st century plant biotechnology. Plant Biotechnol J 2005;3:141-55. http://dx.doi.org/10.1111/j.1467-7652.2004.00113.x.         [ Links ]

73. Newhouse K, Singh B, Shaner D, Stidham M. Mutations in corn (Zea mays L.) conferring resistance to imidazolinone herbicides. Theor Appl Genet 1991;83: 65-70. http://dx.doi.org/10.1007/BF00229227.         [ Links ]


*Corresponding author: E-mail address: rriegel@uach.cl (R. Riegel).

Received 17 March 2015, Accepted 30 July 2015, Available online 9 September 2015

Copyright © 2015 Pontificia Universidad Católica de Valparaíso. Production and hosting by Elsevier B.V. All rights reserved. Peer review under responsibility of Pontificia Universidad Católica de Valparaíso.

 

Creative Commons License Todo el contenido de esta revista, excepto dónde está identificado, está bajo una Licencia Creative Commons