plants
Review
A Review of Chenopodium quinoa (Willd.) Diseases—An
Updated Perspective
Carla Colque-Little, Daniel Buchvaldt Amby and Christian Andreasen *
Department of Plant and Environmental Sciences, University of Copenhagen, Højbakkegaard Allé 13,
DK2630 Taastrup, Denmark; cxl@plen.ku.dk (C.C.-L.); amby@plen.ku.dk (D.B.A.)
* Correspondence: can@plen.ku.dk; Tel.: +45-51322551
Citation: Colque-Little, C.; Amby,
D.B.; Andreasen, C. A Review of
Chenopodium quinoa (Willd.)
Diseases—An Updated Perspective.
Abstract: The journey of the Andean crop quinoa (Chenopodium quinoa Willd.) to unfamiliar environments and the combination of higher temperatures, sudden changes in weather, intense precipitation,
and reduced water in the soil has increased the risk of observing new and emerging diseases associated with this crop. Several diseases of quinoa have been reported in the last decade. These include
Ascochyta caulina, Cercospora cf. chenopodii, Colletotrichum nigrum, C. truncatum, and Pseudomonas
syringae. The taxonomy of other diseases remains unclear or is characterized primarily at the genus
level. Symptoms, microscopy, and pathogenicity, supported by molecular tools, constitute accurate
plant disease diagnostics in the 21st century. Scientists and farmers will benefit from an update
on the phytopathological research regarding a crop that has been neglected for many years. This
review aims to compile the existing information and make accurate associations between specific
symptoms and causal agents of disease. In addition, we place an emphasis on downy mildew and its
phenotyping, as it continues to be the most economically important and studied disease affecting
quinoa worldwide. The information herein will allow for the appropriate execution of breeding
programs and control measures.
Keywords: causal agents; downy mildew; pathogenicity; Peronospora; resistance factors; severity;
quinoa diseases; quinoa disease assessment
Plants 2021, 10, 1228. https://
doi.org/10.3390/plants10061228
Academic Editors: Cataldo Pulvento
and Didier Bazile
Received: 9 April 2021
Accepted: 7 June 2021
Published: 16 June 2021
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Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
1. Introduction
Agriculture is affected by global climate change. Non-traditional crops with high
nutritional value and the ability to cope with abiotic stress are of special interest in today’s
world. Quinoa (Chenopodium quinoa Willd.) is an ancient crop that exhibits remarkable
tolerance to frost, salt, and drought. Moreover, it is highly nutritious and has a vast genetic
diversity resulting from its fragmented and localized production over the Andean region.
The recent introduction and cultivation of quinoa in novel environments has resulted in
a wider spectrum and higher intensity of infectious diseases. Oomycetes and fungi are
the two most important eukaryotic plant pathogens [1]; their predominance on the quinoa
pathobiome is also evident.
Diseases of quinoa have been reviewed previously [2–6]. However, an update is
necessary because new emerging diseases of the quinoa mycobiome are being discovered. Taxonomy based on the morphological characteristics and nomenclature of fungi
is relatively conserved and informative when high-level classifications (genus level) are
considered. However, there is uncertainty when lower-level phylogenies (species level) are
considered due to the fast-evolving traits and phenotypic plasticity of fungi [7]. As a result,
DNA and molecular sequence-database comparisons techniques have been employed,
along with various DNA fingerprinting and more advanced and complex methods such as
whole-genome sequencing, for the identification of plant pathogens [8,9].
The universal nuclear ribosomal primers developed by White et al. (1990) for PCR
amplification of the internal transcribed spacer (ITS) region have become a key component
Plants 2021, 10, 1228. https://doi.org/10.3390/plants10061228
https://www.mdpi.com/journal/plants
Plants 2021, 10, 1228
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the description and characterization of fungal diversity [10]. In addition to ITS, various other
markers exist for multi-locus sequencing. It is commonly used by combining ITS with other
relevant genomic regions (e.g., COX I, calmodulin, and TEF1 gene regions). It has proven
helpful and necessary for the accurate identification of microbial plant pathogens [11–13].
Such molecular approaches should be paired with pathogenicity assessments, including the
description of disease symptoms, isolation and artificial inoculation of quinoa tissue, recording
of symptoms, and re-isolation. These tests are known as Koch’s postulates [14–16]; their
validation discriminates an opportunistic association from a pathogenic-type interaction.
This review aims to provide an updated overview of microbial plant pathogens
causing disease in quinoa, focusing on the morphological characterization and molecular
identification of the causal agents. Research carried out in the Andean countries some
decades ago provides insightful and valuable reports, described herein. We compiled and
analyzed existing information, with a marked emphasis on downy mildew.
2. Downy Mildew of Quinoa
2.1. Nomenclature and Distribution
The oomycete Peronospora variabilis Gäum. 1919 [17] is the causal agent for downy
mildew on C. quinoa (www.indexfungorum.org, accessed on 10 June 2021) and C. album L.
The genus Peronospora belongs to the Peronosporaceae family (Peronosporales order), which
are highly physiologically specialized, biotrophic organisms. Phytopathogenic oomycetes
are eukaryotic microbes with filamentous vegetative growth and spores for reproduction
(fungus-like). Molecular analysis revealed they are among the Stramenopiles (or heterokont),
closely related to golden-brown algae and diatoms [1,18–20]. Fundamental features are:
1.
2.
3.
Oomycetes cell walls are mostly composed of glucans, in contrast to chitin from
fungi [14].
Most oomycetes are insensitive to azole fungicides (e.g., ketoconazole) because they
do not have the ergosterol pathway needed to activate the azole-fungicide mode of
action [21–23].
During their vegetative state, oomycetes are diploid compared to haploid or dikaryotic
fungi [1].
Due to taxonomic confusion, downy mildew was previously classified as Peronospora
farinosa and considered as such by most studies for about 50 years [24–26]. Byford
(1967a,b) [27,28] investigated cross-inoculation experiments and concluded the division of
three formae speciales (f. spp.) Table 1.
Later, a phylogenetic study on P. variabilis of C. quinoa and C. album from different
geographical regions showed that both are located in the same phylogenetic cluster with
no evidence to separate them into different taxa [29–32]. Morphological, molecular, and
biological host specialization analyses revealed that a narrow species concept is more
appropriate for the downy mildews. The available evidence strongly suggests that the
host range of P. variabilis is limited to C. quinoa and C. album [29], that of P. effusa is limited
to spinach [33,34], and more recently that of P. chenopodii has been shown to be limited
to C. hybridum L. (maple leave goosefoot), P. chenopodii-ambrosioides to C. ambrosioides L.
(Jesuit’s tea, Payqu), P. chenopodii-ficifolii to C. ficifolium Sm. (fig leave goosefoot) [13], P.
chenopodii-polyspermi to C. polyspermum L. (many-seeded goosefoot), and P. schachtii to sugar
beet [26]. In older literature, P. farinosa was used as the causal agent of downy mildew of
quinoa. However, the species name “farinosa” had been ascribed to an unrelated genus
(Atriplex) and is no longer valid as a species name for Peronospora [35].
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Table 1. Peronospora species current identity and classification by Byford [27,28].
Plants 2021, 10, x FOR PEER REVIEW
√
√
√
√
√
Host (Genus/Species)
Pathogen
Current Identity
Byford Classification (f. spp.)
P. farinosa formae speciales
Beta spp.
C. álbum + C. quinoa
Spinacia oleracea
P. schachtii [26]
P. variabilis [29,30]
P. effusa [33,34]
P. farinosa f. sp. betae
P. farinosa f. sp. chenopodii
P. farinosa f. sp. spinaciae
The earliest report of downy mildew infecting quinoa in South America came
from Martin Cardenas (1941), who found it infecting quinoa in Cochabamba, Bolivia,
and identified it as P. farinosa [36]. P. variabilis has been documented throughout the
world (Figure 1) [26,29–32,37–52] wherever quinoa is cultivated. It is expected to become
ubiquitous in all quinoa cropping areas as oospores found in seeds have also been seen
in old dried leaves [32,53,54]. Moreover, C. album (known as goosefoot, fat hen, or lamb’s
quarters) [55] is frequently infected by downy mildew throughout Europe because it is
conspecific [56] with the P. variabilis from C. quinoa. Therefore, it is likely to be a reservoir
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for the pathogen and an alternative host [29,52,56]. Other Chenopodium species, such as
C. murale L. (nettle leaf goosefoot), C. ambrosoides L. (Indian goosefoot, Mexican tea), C.
berlandieri Moq. (pit seed goosefoot), and C. ficifolium Sm. (fig leaf goosefoot), were reported
√
√the pathogen based on morphological identification [39,45,57] and molecular
to harbor
COX2 bar coding for C. berlandieri var macrocalycium (Table 2). These reports require
√
√
further investigation to confirm the accurate identity of the pathogen. Cross-infection
reported
√ so far is solely that of P. variabilis isolated from C. album and C. quinoa [52].
Figure 1. Downy mildew disease of Chenopodium spp. distribution map (CAB international, last
modified 21 November 2019 via www.cabi.org/isc/datasheet/39704 (accessed on 10 June 2021).
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Table 2. Documented reports for downy mildew on C. quinoa and weedy Chenopods.
Country
C. quinoa
√
Leaves ( ), Seed (x)
Mor.
Mol.
C.album
Leaves
Mor.
Mol.
C. berlandieri
var. Macrocalycium
Mor.
Mol.
C. murale
Leaves
Mor.
Mol.
C. ambrosoides
Leaves
Mor.
Mol.
C. ficifolium
Leaves
Mor.
Researcher
Year
[Ref]
Mol.
Bolivia
√
Martin
Cardenas
1941
[36]
Peru
√
G. Garcia
1947
[37]
Canada
√
JF.Tewari
1990
[38]
Peru
√
L.Aragon
1992
[39]
Ecuador
√
Jose Ochoa
1999
[40]
Denmark
√
S. Danielsen
2002
[41]
Poland
√
Panka
2004
[42]
India
√
A. Kumar
2006
[43]
Bolivia
√
Erica Swenson
2006
[44]
2008
2010
[26]
[29]
S. Danielsen
2010
[30]
√
√
√
√
√
√
√
√
√
√
√
Argentina
China
Ireland
South Korea
Netherlands
Germany
Latvia
Romania
Italy
√
Y.J. Choi
√
India
P. Baisvar
2010
[45]
√
√
Ana Testen
2012
[46]
Ana Testen
2014
[32]
√
x
x
x
√
Y.J. Choi
2014
[47]
Bolivia
Ecuador
USA
Korea
√
√
√
√
√
√
√
√
√
√
√
√
Peru
Ecuador
Denmark
USA (Pennsylvania)
√
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Table 2. Cont.
Country
Morocco
Egypt
USA (N.
Hampshire)
C. quinoa
√
Leaves ( ), Seed (x)
Mor.
Mol.
√
√
√
√
Turkey
Turkey
√
√
Denmark
√
√
C.album
Leaves
Mor.
Mol.
√
√
√
√
√
√
C. berlandieri
var. Macrocalycium
Mor.
√
x *
Mol.
C. murale
Leaves
Mor.
Mol.
C. ambrosoides
Leaves
Mor.
Mol.
√
C. ficifolium
Leaves
Mor.
√
x *
√
Researcher
Year
[Ref]
Manal Mhada
2014
[48]
Walaa Khalifa
2018
[49]
Helen Nolen
2019
[50]
M.Kara
2020
[31]
Esra Gül
2021
[51]
C. Colque-Little
2021
[52]
Mol.
Mor. = morphological characterization; Mol. = molecular identification. Source: elaborated from references on the column [Ref]. x * Koch postulates failed;
**
√
** corresponds to a field population.
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2.2. Infection Biology and Disease Symptoms
Based on various scientific studies, we assembled a hypothetical disease cycle for P.
variabilis (Figure 2).
Figure 2. Proposed disease cycle of quinoa downy mildew caused by Peronospora variabilis (Photos: C. Colque-Little). Picture
of haploid gamets adapted from Judelson [58].
When mature sporangia fall on compatible leaf tissue with free moisture and relative humidity (more than 85%), the infection begins. Spores from pathogenic oomycetes
produce an adhesive vesicle on the spore side in contact with the host (ventral) at early
infection stages (Figure 3F). Next, a germ tube that faces the host is produced and grows
chemotropically toward a suitable penetration site. In most downy mildews, the hyphae
enter the leaf via stomatal pores [58] (Figure 3C). The formation of an appressorium-like
swelling (penetration structures that exert pressure) on histopathological samples was
observed under a microscope [52,59]. It penetrated the stomata (Figure 3B,D) but did
not directly penetrate the cuticle [49,59]. Spores are chemotaxically and mechanically
dependent on the stomatal aperture [60,61].
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Figure 3. Quinoa leaf infections caused by P. variabilis sporangiogenesis during the early stages of asexual reproduction.
(A) Sporangium forming germ tube (gt) and faint penetration hyphae towards the mesophyll. (B) Extracellular matrices (em)
secreted from germinating sporangium (sp) and appressorium-like (als) structure penetrating stomata. (C) Sporangium,
forming germ-tube (gt) and appressorium like structure in water. (D) Sporangiophore (spr) emerging from stomata.
(E) Sporangiophore holding sporangia, emerging from lower epidermis. Scale bar: 20 µm. (F) Hypothetical P. variabilis
sporangiogenesis timeline (Photos: C. Colque-Little). Illustration in timeline created with Biorender.com.
Stomata colonization happens relatively quickly. Once an appressorium is established,
the secretion of extracellular matrices during the germination of the sporangia appears, as
reported elsewhere (Figure 3B) [59]. The hyphae ramify intercellularly, forming haustoria
(feeding structures) through the leaf tissue five to six days after penetration (Figure 3A–F). The
sporangiophores emerge from the leaf’s surface around the seventh day, carrying asexual
lemon-shaped sporangia (Figure 3E,F). Seven to ten days after the primary infection,
sporangia are disseminated to other leaves by wind and water [40,59] (Figure 3F). They
are assumed to be of importance for spreading the disease during the growing season at
this stage [62]. In general, Peronospora species require moderate temperatures (10 ◦ C–20 ◦ C)
for optimal sporulation [63,64]. While the disease is developing, several asexual cycles
(reproduction of sporangia) may occur. Secondary infection demonstrated that the disease
could spread rapidly in the field if the optimal conditions are present [54].
Infected leaf tissue manifests lesions and signs on both sides of the leaf. Sporulation
becomes apparent mostly on the leaf surface. Symptoms on infected plants vary depend-
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ing on genotype, growth stage, and environmental conditions (Figure 4A−D). Classic
symptoms include pale or yellow chlorotic lesions on the leaf surface (Figure 3E) and dark
gray-violaceous sporulating areas, mostly on the lower surface (Figure 3F). The lesions can
be several and small in some cultivars, whereas in others the lesions are extensive, diffuse,
and irregular (Figure 4G). Lesions turn pink, red, purple, or light-brown, depending on the
plant’s pigments (red-violet and yellow betalains [65,66]). A hypersensitive response has
also been observed (Figure 4E,G). The sporulation presence differs considerably, probably
due to cultivar responses and the pathogenic capability of the specific isolate [52,54,67].
Figure 4. (A) Quinoa crop severely damaged by downy mildew. (B–D). Infected varieties in the
fields of the main quinoa growing areas of Bolivia. (E) Adaxial leaf side belonging to different
quinoa genotypes artificially infected with downy mildew. (F) Abaxial side of the leaves showing
sporulation. (G) Differences in disease symptoms, ranging from hypersensitive reactions causing
pale yellowish spots (left) to high susceptibility with chlorotic lesions covering the whole leaf (right)
(Photos: C. Colque-Little).
Downy mildew primarily affects the foliage, but it is possible to find it colonizing
different organs and tissues of quinoa plants. However, its symptoms are less obvious and
sporulation is inexistent. Therefore, polymerase chain reaction (PCR) was used to amplify
P. variabilis DNA. Taha (2019) gathered a composite of quinoa seedlings at different growth
stages, subdivided them into different organs, and detected P. variabilis DNA on 0.8% of
the root samples, 83% on the cotyledon and leaf, and 42% on steam samples. The PCR was
also positive for 60–80-day-old plants’ inflorescences [68]. In addition, scanning electron
microscopy was capable of visualizing P. variabilis on petioles [59], and the mycelium was
seen as in the intercellular spaces of the leaf midrib of 80-day old plants [69]. Since the
Plants 2021, 10, 1228
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pathogen was detected at early and late growth stages of the quinoa plant, it was thought
to present a systemic mode of infection [68]. However, other researchers argue [69] that the
germinated oospores-mycelium spreads through intercellular parenquimatic spaces (next
to xylem but not wood vessels) of the hypocotyl acropetally, towards the plant’s aerial
parts, and is finally inserted into the developing seed. For clarification of the mode of
infection of P. variabilis, more research is needed.
2.3. Morphology and Reproduction
Peronospora variabilis hyphae are coenocytic (hyphae without septae) and multinucleate, resulting from nuclear divisions within the cell without an accompanying division of
the cytoplasm (cytokinesis). Sporangiophores are 240−580 µm long, slender, arborescent,
dichotomously ramified five to six times in a sharp angle, ending in two to three straight to
slightly curved branches (Figure 5A). Ultimate branchlets are in pairs or single, flexuous to
curved 8−23 (av. 12.3) µm long, with obtuse tips (Figure 5C) [29]. Sporangia are pedicellate,
deciduous, olivaceous with a grayish tint, broadly ellipsoidal to ellipsoidal (av. 27.7 µm
long × 21.0 µm wide (Table 3), ending in an apical translucid papilla [47]. Taxonomic
measurements such as spore lengths and widths can vary depending on the homogeneity
of the conidium population, the origin of the isolates, the spore subpopulation, or different
roles or times in the pathogen’s life history [70]. Measuring that variability under the
microscope allows researchers to estimate the mean length/width with a reasonable level
of resolution when a minimum of 41–71 spores are measured for the Peronospora genus [71].
Even though P. variabilis, infecting C. quinoa and C. album, is conspecific [56], sporangia
found in C. album were slightly bigger. Further research is needed to figure out why this
difference exists. Table 3 illustrates this variability from measurements taken by various
researchers [26,29,31,46,47,49,51,54,72] and the average of their measurements is provided
as a reference (Figure 5B,D).
Table 3. Peronospora variabilis sporangium sizes when isolated from C. quinoa and C. album.
P. variabilis Sporangium Isolation Origin
C. quinoa
av. Length × Width (µm)
25.5 × 17.5
22 × 23.13
27.5 × 20
28.8 × 21.8
30.7 × 23.8
31 × 23
28.5 × 23.5
27.7 × 21.0
1.32
C. album
av. Length × Width (µm)
29.5 × 23
30 × 25
30.1 × 24
1.25
Reference
Khalifa and Thabet 2018 [49]
Yin et al., 2018 [72]
Gül, 2021b [51]
Danielsen & Ames, 2004 [54]
Choi et al., 2010 [29]
Testen et al., 2012 [46]
Choi et al., 2014 [47]
Choi et al., 2008 [26]
Kara et al., 2020 [31]
av. size
av.ratio
Peronospora variabilis can reproduce asexually (sporangiogenesis) and sexually (oospore
formation and germination). It has been reported to be heterothallic and requires two
compatible partners for oospore formation (mating). When eight single-lesion isolates
coming from different regions of Peru and Bolivia were crossed in all possible combinations
using a detached leaf assay, the existence of two mating types, P1 and P2, was apparent [73].
Sexual cycles start with a male (antheridium) and a female (oogonia) gametangia. These
structures can be observed in the leaf mesophyll of plants sown 45 days earlier [69] and
have the appearance of swollen hyphal tips [74]. Once in contact, both swell, especially
the oogonia. Next, synchronous meiosis occurs within each one, and a pore develops
between them. A single haploid nucleus is then transmitted from a male to a female. After
fecundation, the development of an oospore starts by establishing a thick multi-layered
Plants 2021, 10, 1228
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wall. During maturation, the ribosomes and cytochromes disappear. The combination of
their lowered metabolism, thick wall, and lipid-rich cytoplasm make them effective resting
structures. Walls are usually hyaline, yet contain a brownish pigment, and their thickness
ranges between 3 and 6 µm in most Peronosporales [58]. Peronospora variabilis oogonia
(isolated from C. album) are subglobose with an average diameter of 43.5 µm [26,31]. The
oospore shape is globose to ovoid; their color varies from transparent to golden brown to
brown [53,75].
Ooospore diameter has been reported to range from 18.2 to 44.5 µm on average [53,69,75]
when isolated from C. quinoa, compared to 25 to 44.5 µm on average when isolated from C.
album [26,54]. These differences may be due to interactions with the host, environmental
conditions, the age of the spore, or the pathogen races [53] (Figure 5 Aa,D). Oospores can
survive inhospitable environments, such as freezing, desiccation, starvation, and microbial
degradation [19]. They permit the completion of the pathogen life cycle and enhance its
fitness by providing a mechanism for genetic variation [58]. Resting structures are often
the source of initial infection. El-Assiuty (b) et al. (2019) hypothesize that oospores bearing
tissues (cotyledons, leaves, and the perianths of seeds) shed during the life cycle of quinoa
plants may play a role in the persistence of oospores in soil.
Danielsen and Ames (2004) [54] detected oospores in the pericarp (external tegument
of the episperm) using ultra-microtome cuts (Figure 5F). El-Assiuty (a) et al. (2019) confirmed their occurrence in examined seed samples, revealing a 90% presence in the perianth,
87% in the seed coat, 3% in the embryo, and 2% in the perisperm [53].
To follow the passage of P. variabilis inside tissues, El-Assiuty (b) et al. (2019) conducted histopathological/microscopic investigations at different plant growth stages. After
planting the surface-sterilized seed of a downy mildew susceptible variety, the observations
started. Oospores were present in the radicle-pith three days after germination, inside
the cortex of hypocotyls, and in the mesophyll of cotyledons seven days after planting.
Oospore germination started with two undulating germ-tubes located opposite to one
another. They develop in the cortex tissue of juvenile seedlings 15 days post-planting [69].
This research is consistent with what has been found for other downy mildew diseases
such as Plasmopara viticola, the mature oospores of which germinated for 3−7 days under a
favorable regime of rainfall and temperature [76].
Moreover, oospores were detected in all tissues of quinoa plants that had been sown
45–120 days previously [53,69]. They were also seen on leaves of senescent infected plants,
artificially inoculated with a single Danish isolate under greenhouse conditions, suggesting
that the isolate had both mating types present (Figure 5E. Colque-Little, unpublished). In
addition, they have been found in old infected leaves collected in Andean regions of quinoa
production (Peru, Bolivia) [54] and fresh leaf tissue collected in Pennsylvania, USA [77].
Greenhouse experiments with oospore-infected seed samples sown in high and low
relative humidity showed a significant difference in visible seedling infections among
samples under high humidity and with a large oospore density in most cases. However,
oospore density seems to be more critical for seedling infections when the relative humidity
is low [75,78,79]. The number of oospores can be estimated using the seed washing
method [54,80,81]. Briefly, the seed is soaked in water under agitation. Seeds are removed
with cheesecloth, the solution centrifuged, the supernatant discarded, and the pellet is
dissolved in sterile water. The number of oospores is counted using a hemocytometer
under the microscope. Calixtro (2017) quantified the number of oospores present on
susceptible seeds and found it was three times greater than the number on tolerant varieties
demonstrating that host genotype is an important factor [82].
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Figure 5. Peronospora variabilis spores isolated from C. quinoa. (A) Sporangiophore with lemon-shaped
sporangia. (Aa) Oospores–sporangia size comparison. (B) Oospore on top of dried leaf tissue. Scale
bars: 20 µm (Photos: C. Colque-Little). (C) Schematic representation of oospore localization in quinoa
seed. o = oospore; pe = pericarp; en = endosperm; p = perisperm (illustration adapted from Danielsen
and Ames (2004) [54] and Prego et al. (1998) [78]).
2.4. Peronospora variabilis Genotypic Diversity and Virulence Profiling
Peronospora variabilis is a genetically diverse group [30] with multiple population
structures, in light of three facts:
1.
2.
3.
Chenopod hosts have a vast degree of genetic diversity and plasticity [83,84].
Peronospora variabilis has great adaptability (climatically and geographically), hence
its worldwide geographic presence [54].
The occurrence of sexual reproduction permits genotypic pathotype expansion [4].
Quinoa cultivation areas of the Andean region have resulted in severe infections under field conditions. Swenson (2006) collected 43 isolates from eight Bolivian regions [44].
Phylogenetic fingerprinting relationships revealed high genotypic diversity within a geographical region. The most recent fungal and oomycete identification initiatives were
carried out using DNA sequencing [12]. A group of P. variabilis herbarium and isolates
from different geographic locations (Argentina, Bolivia, Denmark, Ecuador, and Peru) were
phylogenetically analyzed based on ITS rDNA sequences. The majority of the Danish
and South American isolates were separated into two major clusters [29]. P.variabilis was
detected in 31 out of 33 quinoa seed lots destined for human consumption and originated in six different countries. Subsequently, ITS and Cox2 phylogenetic relationships
were examined to determine whether geographical differences occurred. ITS-derived
phylogeny showed no genetic differences, but the Cox2 phylogeny indicated that geographical differences existed between US and South American samples [32]. In another
study, researchers characterized 40 isolates from P. variabilis originating in the Andean
highlands (Peru and Ecuador) and Denmark (Jutland, Sealand) using universally primed
PCR (UP-PCR) fingerprinting analysis. A separation between the Danish and Andean
isolates in two distinctive clusters was found, together with genotypic variations between
isolates within each cluster [30].
In the future, the next step might be the virulence profiling of P. variabilis, achieved
through the sequencing of its genome, followed by transcriptomic analysis. Progress in
genome sequencing technologies can provide genome data to better understand how microbes live, evolve, and adapt. Indeed, the genome of three races of P. effusa (downy mildew
of spinach) was recently sequenced, assembled, and annotated to gain insights into its gene
repertoire and identify infection-related genes [9]. The genomes of microbial pathogens
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can vary greatly in size and composition; this also includes when closely related species
are considered. In the case of Peronospora, species greatly vary between 45.6 to 159.9 Mb
when estimates are made using image analysis of nuclear Feulgen staining [85]. Whether
genome sizes have an impact on the lifestyle of Peronospora species is still unknown [86].
Another way to elucidate genotypic and phenotypic variation within pathogen populations is to use virulence-phenotypic assays with a standard set of differential hosts.
Spinach downy mildew has such a set composed of 11 cultivars, maintained with the help
of the international working group on P. effusa (IWGP) [86–88].
This organization invites researchers to use the set to identify new isolates that can
later be nominated, tested for various criteria, and then given a race designation [86–88]. An
international system for monitoring the virulence of P. variabilis has not yet been developed.
However, Ochoa et al. (1999) made the first step towards this from a collection of twenty
P. variabilis isolates that corresponded to different Ecuatorian ecoregions [40]:
1.
2.
3.
An area where quinoa cultivation was not regularly practised. The least virulent
strains were present here and were identified as virulence group 2 (V2).
A region where landraces and newly released cultivars were introduced. Only the
most virulent strains belonging to group 4 (V4) were present here.
Fields located where landraces and newly released cultivars have been cultivated for
many years. Here, all four virulence groups were present.
Ochoa et al. (1999) investigated seedlings under controlled environments from 60
selected genotypes and the above-mentioned P. variabilis collection; quinoa lines were
selected for consistent compatible/incompatible reactions. Based on these results, four
resistance factors (R1, R2, R3, and R4) were postulated [40]. It was most likely that two
mating types are present. areasall difference exists. However, these genotypes are exclusive
to the National Ecuadorian collection and thus not available for research. The measurements
of severity and sporulation of downy mildew from reference cultivars (Puno, Titicaca, and
Vikinga) and many other genotypes used by Colque-Little et al. (2021) are comparable
to the 1−5 scale developed by Ochoa et al. (1999) (Table 4 and Figure 6). Therefore, we
suggest that the presence of resistance factors could be preliminarily hypothesized on
reference cultivars. Importantly, the seed of these cultivars is commercially available
(Quinoaquality.com, Denmark) and could be established as an international reference set.
Table 4. Set of quinoa cultivars and Chenopodium album postulated for profiling the virulence of Peronospora variabilis.
Cultivar
Hypothesized
Resistance Factors
C. album
Puno
Rosa Blanca
Blanca
Titicaca
Vikinga
R1, R2, R3, R4
R1, R2, R3, R4
R1, R2, R3
R1, R2, R3
R1, R2
R1
Response to Downy Mildew
% Severity
% Sporulation
5
11
32
46
52
70
0.04
0.2
17
47
40
69
Origin
Denmark
Denmark
Bolivia
Bolivia
Denmark
Denmark
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Figure 6. Set of reference cultivars postulated for profiling the virulence of Peronospora variabilis, including C. album and two
Bolivian cultivars with intermediate reactions. Leaves from three-week-old artificially inoculated plants. Numbers in red
indicate the percentage of severity on the adaxial side, and those in purple indicate the percentage of sporulation on the
abaxial side [52]. Numbers in green (incompatible response) and blue (compatible response) correspond to Ochoa’s scale:
0 = no symptoms; 1 = 2−5-mm lesion with truncated mycelium in the mesophyll of the leaf; 2 = 4−8-mm chlorotic lesions
with minor sporulation; 3 = medium-sized and confined chlorotic lesions with sporulation mainly on the abaxial side of
the leaf; 4 = large, not clearly confined chlorotic lesions with sporulation mainly on the abaxial side of the leaf; 5 = mild
chlorosis with abundant sporulation on both adaxial and abaxial sides of the leaf (Ochoa et al., 1999 [40], Colque-Little et al.,
2021 [52]). Both assessments are comparable in terms of severity and sporulation; thus, the existence of resistance factors is
hypothesized in this set of reference cultivars.
2.5. Disease Assessment under Controlled Conditions and in the Field
Reliable identification, followed by the assessment of disease, is the first step in
efficient management. It is also an important component in the development of diseasetolerant quinoa varieties. It allows for crop-loss assessments and screening for host–
pathogen interactions. Assessment methods must be in close agreement with the goals of
the trial(s). Evaluations might differ according to the experimental setup. For seedlings,
detached leaves and plantlets under controlled conditions and a disease assessment scale
can be used (Figure 6). For the assessment of diseased plants in the field, it is necessary to
take into account:
1.
The phenological stage of the plants. Age-related resistance becomes relevant for
biotrophic pathogens, which require healthy plant tissue to complete their cycle. [89,90].
The observation of symptoms should reflect the progression of the disease through
periodical records, rather than observing its percentage of occurrence or incidence.
For the quinoa/downy mildew interaction, it has been demonstrated that disease
incidence has a low heritability H2 = 0.4 and a low correlation with severity and
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sporulation (0.67 and 0.65, respectively) [52]. Therefore, incidence or whole plant
scores are unsuitable for this type of trial. To measure the area under the disease curve
progression (AUDPC), a minimum record of three to four observations of disease
severity is essential. A similar study has highlighted the importance of measuring
the disease severity over time for other interactions, such as Phytophthora infestans
infecting potatoes. The objective is to capture low, medium, and high infection levels
in all the genotypes, including the susceptible ones [91].
Calixtro (2017) recorded high variability in the area under the disease progress curve
(AUDPC) within the same quinoa accession during different phenological stages. The
higher AUDPC values were seen at 104 days after sowing with favorable disease conditions [82].
Therefore, we suggest assessing downy mildew as soon as the first symptoms of the
disease are visible. The first reading could be when nine pairs of leaves (BBCH 1–1.9) have
emerged or beforehand in cases where disease symptoms are visually observed. Time
intervals among subsequent readings depend on whether the disease advances slowly
or quickly [91]. Other observation points could be during development (BBCH = 4) or
visible inflorescence (BBCH = 5–5.9) and the last one at complete anthesis (BBCH 6–6.9).
The phenological growth stages mentioned here correspond to the international quinoabased coding system BBCH (Biologische Bundesanstalt Bundessortenamt und Chemische
Industrie) [92].
2.
The vegetative cycle of the plants. Late-maturing quinoa genotypes will display some
degree of resistance [93] by increasing the latent period of the pathogen. Thus, readings for
severity were taken ten days after infection instead of five in a recent study, in which a late
cultivar Blanca was compared with the Danish cultivars Puno, Titicaca, and Vikinga [52]
(Figure 7). Puno matures ten days later than Titicaca [94]. Cultivar Blanca is considered
susceptible when additional time is given [52] (Figure 7). Vegetative cycle effects were
also shown in another study that analyzed the mean-based cluster of inter-ecotype
F2:6 population crosses and identified the following three clusters [48,95,96]:
(a)
(b)
(c)
Cluster one: consisting of late, mildew-resistant, high-yielding lines;
Cluster two: consisting of semi-late lines with intermediate yield and
mildew susceptibility;
Cluster three: consisting of early to semi-late accessions with low yield and
mildew susceptibility.
Therefore, for a proper comparison, quinoa lines with similar vegetative cycles should
be screened in the same experiment or statistical adjustments should be carried out as part
of the analysis. In addition, a positive control (susceptible variety) and a negative control
(resistant variety) might be beneficial in the analysis.
3.
Sampling method and sample size. Depending on the size of the experiment, there is
no need to take severity readings in all the quinoa plants. Instead, consider the plot
level and take readings on representative samples. Normally, 6−10 plants per plot are
sufficient [54,91]. Next, an estimation of the percentage of affected foliage is required.
Given the size of the plants and abundant foliage, it is not feasible to analyze the entire
foliage; thus, it is recommended to perform scoring on individual leaves of the chosen
plants [54]. Danielsen and Munk [97] evaluated various field assessment methods to
predict yield losses due to downy mildew. The three-leaf method resulted in the highest
negative correlation to yield (r = −0.736). Furthermore, disease progression relies on the
successful infection of the host. It is often assumed that the susceptibility of host tissue
is constant. However, in reality, it is a function of plant age and leaf position [14,98,99].
These responses might result from inducible plant defense responses, which occurs
at the starting interaction site but also in distal, uninfected parts [99–101]. For these
reasons, we suggest randomly choosing three leaves from the middle part (lower third,
middle third, and upper third), as illustrated in Figure 8. Avoid the lower and upper
extremities of the plant because they are prone to senescence/defoliation [97] and plant
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defense responses, respectively. Next, estimate the percentage of affected leaf area using
the attached scale [79] (Figure 9). The average value from the score of the three leaves
becomes the percentage of severity for each plant.
Figure 7. Reference cultivars’ responses to infection with Peronospora variabilis, measured in mean
severity under greenhouse conditions. Source: Colque-Little et al. [52].
2.6. Yield Losses and Management
The losses caused by downy mildew depend on the plant’s phenological phase at
the time of infection and the amount of resistance that the cultivar has [102]. Infection
of susceptible cultivars may result in severe yields losses if the pathogen has favorable
weather conditions, particularly high relative humidity [54]. If the infection occurs in the
plant’s initial growth stages, susceptible crops could completely fail; in less susceptible
cultivars, the loss may fluctuate between 20% and 40% [4]. In a conventional intensive
agriculture system of Cajamarca (Peru), between five to seven fungicide applications were
needed to control the infection during the agricultural campaign [103].
Due to the high capability of P. variabilis for the proliferation and latent infection
on C. quinoa and C. album, the scenario for low-input farming has only two options for
disease control:
1.
2.
tolerant crop varieties; and
cultural practices (options on the list below).
Alternative cultural practices:
(a)
(b)
(c)
Policymakers, smallholder farmers, and other stakeholders need resources for collective action for the establishment of a seed supply chain with quality standards (low
levels of key seed-borne diseases). Experiences with complementary intervention
such as capacity building and technical assistance have shown this influence in an
appropriate conceptual model of sustainable production [104].
The detection of P. variabilis on the seed is achieved using a simple method [32,54]. In
the case of the presence of an oospore, treat the seed with a systemic fungicide [105].
For small samples, alternative treatments such as a hot water bath (50 ◦ C–60 ◦ C)
could be considered for 10 to 30 min, as this method has been applied successfully to
eradicate seed-borne pathogens of spinach [106]. After or without treatment, the addition of beneficial microbes by priming the seed with products such as commercially
available Trichoderma can enhance the growth of the plants [107].
Adjusting the space between rows and individuals, making the area less dense and
increasing space between plants. In areas where the RH is as high as 80%, the
minimum should be a 0.5-m space between rows and 0.15 m between plants [5].
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(d)
(e)
(f)
(g)
Avoiding excess water in the field;
Implementing effective weed control, especially of alternate host C. album;
Practicing crop rotation;
Spraying the plants around 45 days after planting in areas with endemic infection
as a preventive measure [69]. Use oomycete sensitive chemical control measures
(e.g., Alietti) at principal growth stages, e.g., leaf development, inflorescence emergence, flowering, and fruit development [14,54,92]. Fungicides could be applied,
alternating between systemic and contact products, starting with systemic products.
Bio-pesticide or plant extracts could replace fungicides with a uniform and preventive
application [5]. Inducers of resistance are an alternative [108].
Figure 8. Modified from Danielsen and Munk (2004) [97]. Three-leaf field assessment method for
quinoa-downy mildew at different growth stages.
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Figure 9. Scale for percentage of severity and sporulation area affected by downy mildew in quinoa. r = postulated minor genes; R = hypothesized major genes. BOL = accession numbers.
Note: Percentage of sporulation is estimated on the abaxial leaf side area covered by visible lesion. It is not estimated on the total abaxial side leaf area (Colque-Little et al., 2021) [52].
Photos by Colque-Little.
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2.7. Genetics of Resistance to Downy Mildew
For agriculture, field or host resistance is still the most important way of controlling
diseases because it leads to the most cost-effective ratio for the grower [109–111].The response to downy mildew in a diversity panel of 132 quinoa genotypes resulted in strong
phenotypic variation with high disease trait heritability (H2 = 0.78 for severity, H2 = 0.82 for
sporulation). This variability was paired with the analysis of 603,871 SNPs in 61 genotypes
with FarmCPU [52]. A single variant on chromosome 4, located above a threshold with a
lack of siginificant marker trait wide associations. A single variant on chromosome 4, located above a threshold with a lack of significant marker-trait wide associations, suggested
a polygenic architecture for the downy mildew interaction in agreement with other studies [43,48,49,95,104,112–115]. However the interactions of the host resistance pathway with
a biotrophic pathogen (e.g., P. variabilis) are complex. The interaction oscillates between
compatible (susceptible) and incompatible (resistant) states, because the genes involved
can introduce quantitative variations, adding different levels of reactio to the extreme
responses adding different levels of reaction to the extreme responses [116]. The same
study phenotyped hypersensitive responses, most probably corresponding to R-genes,
and very low sporulation on resistant genotypes, which could correspond to defeated
R-genes [52] (Figure 3E,G). Indeed, Gabriel et al. (2012) characterized the quinoa/downy
mildew pathosystem in field experiments and discussed the presence of R-genes, multiple
r genes, defeated R-genes, and combinations, with the most common interaction being
that corresponding to field resistance [93]. The deployment of different genes depends on
many factors, such as pathogen isolate aggressivity [40] and environmental conditions. For
quinoa downy mildew, it was demonstrated that the variance for genotype-by-experiment
interaction σ2 G E was large, reflecting that even minute environmental changes can trigger
a genotype to respond differently to the disease (Figure 10) [52]. The degree of resistance
that the plant displays is determined by these changes interacting with the host genetic
composition [116]. Furthermore, segregation in an F2 mapping population derived from a
cross of saponin-free and bitter genotypes suggested that downy mildew resistance has a
dominant inheritance [117].
Figure 10. Ratio calculated from mean averages of sporulation/severity for the South American
diversity panel. The names inside the histogram bars correspond to reference and representative
cultivars for each group. Source: calculated with the data set from Colque-Little et al. (2021) [52].
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Therefore, field phenotyping experiments of P. variabilis infections using diverse
quinoa genotypes should include multiple environments and points in time. Using mixed
modeling to detect quantitative trait loci (QTLs) by considering them as random samples
from a population of target environments and time could be one alternative [118]. Under
controlled conditions, it would make sense to use elite diversity panels with replicates,
reference cultivars, and genetically diverse pathogen isolates in a series of experiments that
are designed randomly.
The characterization of a south American panel demonstrated robust differences
between the genotypes for all disease traits [52] (Figure 11). Moreover, at least five cultivars
that were released by Bolivian breeding programs [112,119], which included downy mildew
tolerance, showed moderate to low severity and reduced the reproduction of the pathogen.
Interestingly, the incidence (Figure 11A,C) and severity of cultivars 6, 17, and 18 might have
classified them as susceptible, but their ability to prevent the pathogen from multiplying
conferred them some degree of resistance [110] (Table 5). Indeed, the Danish variety Titicaca
was classified as susceptible through the solely detached leaf sporulation assay [48]. When
the assessment was done as a function of both parameters, by calculating the ratio (R =
%sporulation/%severity), Titicaca’s R = (40/52) = 0.77 showed that it is not completely
susceptible (Table 5). This finding suggests that the scoring of both parameters in plantlets
can contribute to better disease assessments of cultivars.
Figure 11. Disease traits estimated means fitted on a generalized linear mixed model (GLMM) for a
diversity panel, comprising gene bank accessions (landraces), cultivars (Bolivian-bred cultivars), and
check varieties (reference cultivars). (A) Severity of infection, (B) sporulation, and (C) incidence of
infection. Error bars represent 95% confidence intervals. Adapted from Colque-Little et al. (2021) [52].
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Table 5. Phenotypic infection traits and Ratio for representative cultivars and reference varieties.
Name
% Severity
% Sporulation
Spo/Sev Ratio
% Incidence
Ratio Based
Classification
C. album
5
0.4
0.08
45
Resistant
Puno
11
0.2
0.02
42
Resistant
Cv6 (Rosa Blanca)
32
17
0.53
59
Highly tolerant
Cv17 (Canchis)
41
30
0.73
73
Mildly resistant
Cv18 (Pandela Roja)
45
29
0.64
74
Mildly resistant
Cv16 (Kurmi)
45
50
1.1
56
Susceptible
Blanca
46
47
1
79
Mildly susceptible
Cv8 (Blanquita)
50
69
1.4
67
Very susceptible
Titicaca
52
40
0.77
81
Mildly resistant
Cv3 (Ayrampu)
52
63
1.2
77
Susceptible
Cv20 (Aynoka)
58
63
1.1
83
Susceptible
Cv21 (Mariqueña)
71
84
1.2
82
Susceptible
Therefore, we propose using the (R = % sporulation/% severity) ratio to better rate
elite genotypes in breeding programs. Using the data set from a previous study [52], the
ratio was calculated. Histograms separated the diversity panel into six groups and derived
a ratio-based scale (Figure 10). The bimodal distribution displayed by the histograms is
consistent with previous findings for P. variabilis field interactions [44].
Quinoa cultivation in South America occurs in agro-climatological polar regions.
These regions have been classified according to their soil type, rainfall, and temperature
as Northern, Central, Southern highlands, and Andean slopes (Table 6) [4,96,112,119].
The Andes have heterogenic topography; their altitude ranges between 3200 and 6500 m
above sea level; hence, there are variations in temperature and humidity [120]. Indeed,
temperature decreases at a rate of 0.7 ◦ C for every 100-m increase in altitude in Chile’s
Tarapaca region. Therefore the coastal Atacama littoral plains differ from mountain sites
(e.g., Los Condores) which enjoy fog oases and lomas vegetation [121]. A similar situation
is expected for the slopes in the Andes of Bolivia, Peru, and Ecuador.
Table 6. Eco-regions for quinoa production in South America.
Eco-Region
Soil
Altitude m.a.s.l
Rainfall (mm)
Max.
Temperature
Min.
Av.
Northern Highland
shores of Lake Titicaca
Rich in organic
matter
3500–4000
500
14
4
7
Central Highland
Slightly acid
3300–4100
350
17.7
8.7
Southern Highland
Arid and poor soils
3200–4000
50–200
18
−2
Andean Slopes
Coastal/Lowland
Northern,
Central, and
Southern
Variable
800–3200
3500–700
12
Variable
Sea level to
Mountain range
40 > 2000
23
21
17
−11
5.7
3
7.6
−8
7
6
4.5
14
11
Within the sub-regions, temperatures vary depending on location (coast or foothills), not shown. Source: elaborated with information
from Gandarillas et al. (2015) [112]; Seiler et al. (2013) [120]; Cereceda et al. (2008) [121]; http://germoplasma.iniaf.gob.bo (accessed on 15
April 2020).
Quinoa ecoregions were inferred from information provided by passport data
(germplasma.iniaf.gob.bo-GRIN global, accessed 15 April, 2020) and the characterization of
Bolivian and Coastal ecoregions [112,120,121]. The information is summarized in Table 5.
Disease traits data (mean values of severity and sporulation) from the South American diversity
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panel [52] were analyzed with the Tuckey test for their relationship with the seed-ecoregion
collection site. For this analysis, we used Inti-Yupana for R [122], and the results pointed at
significant differences for the variables. The graph represents data from means of sporulation
only because data from means of severity was very similar (Figure 12).
Even though the sample size from the central highlands was overrepresented and the
Coastal sample size was underrepresented, significant differences (p = 0.05) for severity
and sporulation were detected. The most resistant genotypes from the South American
diversity panel came from the coastal/lowland and northern highlands ecoregions. The
northern highlands are the most humid since they are close to Lake Titicaca. This ecoregion
is suitable for pathogen infections and disease pressure. This outcome is in agreement
with previous reports [4,48,54,93,95]. The Danish cultivar Puno was found to be resistant,
as reported elsewhere [48]. Moreover, principal component analysis of genome-wide
association studies (GWASs) demonstrated that Puno is genetically close to Chilean coastal
lines and separated from highlander genotypes [52]. The central highlands showed the
largest quantity of susceptible genotypes, likely because they were also the most numerous.
However, a few genotypes with a large amount of sporulation came from the southern
highlands of Bolivia (i.e., G16, G17, and G82) [52] (Figures 11B and 12B).
Figure 12. (A) Modified map of germplasm bank accession across South America by elevation Source: Colque-Little et al.
(2021) [52] and modified map of Bolivian ecoregions for quinoa production. Source: Gandarillas et al. (2015) [6]. (B) Mean
sporulation on diversity panel related to quinoa ecoregions calculated with Tukey test (p = 0.05). Different letters (a,b,c)
represent significant differences between the sporulation produced by genotypes coming from different ecoregions when
infected with P. variabilis.
In conclusion, the genetic improvement of quinoa for downy mildew tolerance is
possible because resistance is present in multiple genotypes, but a virulent pathotype
might overcome it. Other options to consider are discovering, transforming, and deploying
resistant alleles existent in wild species such as C. albums [52,123,124]. Because tolerant
varieties seem to delay and reduce the disease progression, inducers of resistance [125,126]
could be a feasible option [108].
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3. Ascomycete Fungi
3.1. Fungi Identified by Molecular and Morphological Approaches
3.1.1. Ascochyta Leaf Spot and Black Stem (Ascochyta hyalospora and A. chenopodii)
At least two Ascochyta species infect quinoa, causing quinoa leaf spot (described
below) and black stem (described in next section). Quinoa leaf spot is either caused by
Ascochyta hyalospora or A. chenopodii. A. hyalospora Coole and Ellis is an Ascomycete, class
Dothideomycetes, order Pleosporales. It was first found as a seed-borne pathogen of
C. quinoa from the Bolivian central highlands, for which a blotter test (seed incubation
method on well-soaked filter papers [127]) revealed 8%−26% of infection. It was identified
morphologically, followed by pathogenicity tests causing whitish leaf spots 5 dpi, followed
by pycnidia at 10 dpi, and necrosis on leaves and stem of C. quinoa and C. album plants [128].
Testen et al. (2013) isolated a fungal pathogen from quinoa fields in Pennsylvania, USA,
and through DNA sequencing of the ITS1-2 region matched it to Ascochyta sp., and reported
that it resembled the morphological characteristics of A. chenopodii and A. caulina, which
at the time of identification had no DNA bar-codes available for comparison. However,
the ITS1-2 sequences from Testen et al. (2013) were not released as GenBank sequence
data [129]. Thus, it is still not possible to make the comparison.
Ascochyta hyalospora pycnidia are globose to subglobose, usually 17.5 to 25 µm in
diameter [128], and contain sub-hyaline to light-brown-colored conidia. The conidia are
cylindrical to ovoid, measuring 19 × 7.5 µm [129] and 25 × 10 µm [128] on average. They
often have one to two septa and less commonly have three septa. Boerema (1977) noted that
the conidia formed on leaf spots after artificial inoculation were longer (35 µm) and often
had two or three septa (Figure 13E,F). Lesions on the leaves are of irregular shape, and are
bronze to reddish-brown with darker edges. Spots eventually turn necrotic. Thereafter,
numerous black pycnidia, distributed randomly in each lesion, can be seen [129].
Figure 13. Leaves showing symptoms of infection caused by A. hyalospora (A) on the adaxial side of the leaf and (B) on the
abaxial side. (C) Stems showing pycnidia and brown stalk. (D) Stem showing pycnidia. (E) A. hyalospora conidia (Photos:
Testen, 2020) [77]. (F) A. hyalospora: (a) pycnidium (×200); (b) conidiogenous cells of pycnidium (×1000); (c) conidia from
pycnidium (×400); (d) bi and tri-septate conidia from pycnidium on an inoculated stem of C. quinoa (×400); (e) conidia from
pycnidium on leafspot of inoculated leaf of C.quinoa (×400). Source: photos (A−E) provided by A.L. Testen. F. Adapted
from [128].
The stems show necrosis, and the pycnidia are visible to the naked eye (Figure 13A−D).
The seeds turn brown, and pycnidia are observed at the stereomicroscope [128,130]. Ascochyta
leaf spots have been considered of minor importance in the Andean region [3,4,6,131]. In 2014,
large-scale cultivation (12,000 ha) of quinoa started in China [132], where the production
was affected. Infected foliage decays and falls, leaving the plant defoliated [5]. Effects on
quinoa production in the USA have not yet been assessed [129]. Experiments in Bolivia
showed that the germination rates of seeds from infected plants were reduced by 6% to
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10%. Moreover, the disease was transmitted to seedlings [5]. One possibility for control
would be the use of high-quality seeds, since the pathogen is seed-borne [130].
3.1.2. Quinoa Black Stem (Ascochyta caulina)
Molecular and phylogenetic analysis of representative isolates from quinoa blackstem revealed that its causal agent is Ascochyta caulina (van der Aa and van Kesteren
1979). Its sexual teleomorph stage is called Neocamarosporium calvescens (de Gruyter et al.
2009), previously known as Pleospora calvescens. The taxonomic status of P. calvescens
has changed recently, based on multigene analyses. It has been established in the genus
Neocamarosporium Crous and Wingfield in 2014, which comprises 15 species, including
N. betae, N. chenopodii, and N. calvescens. These species share the same large phylogenetic
branch with N. calvescens [133–137]. Ascohyta caulina in its asexual form belongs to the
family Didymellaceae and has often been confused with A. hyalospora [137]. Previously, it
has also been found to infect eight species of Atriplex and eight species of Chenopodium,
including C. album [138].
Another report [139] on A. caulina was accomplished through a morphological description of the isolate found on quinoa seeds of cv. Cochabamba of Bolivian origin (stored
at the Gene Bank of the Research Institute of Crop Production in Prague-Ruzyně). For
pathogenicity tests, the isolate was inoculated in seedlings, and symptoms were reproduced. Interestingly, quinoa seeds from the University of Copenhagen, Denmark, analyzed
simultaneously, were free of A. caulina [139]. This finding might indicate that the disease is
not present in Denmark. A. hyalospora pycnidia are rigid structures, grayish-white or light
brown, spherical or pear-shaped, and have a single chamber. They are 162 × 134 µm in size,
on average. Conidia are elliptical or fusiform, light brown, oblong at the top and flat at the
base, and measure 17 × 6 µm on average [137]. Conidia usually have one septum, which is
erect or curved (Figure 14D). The optimal conditions for its germination are between 15–25
◦ C, RH = 60%. Compared to A. hyalospora leaf spots, black stem lesions were more likely to
develop under cooler conditions [140]. Pathogenicity tests on detached stems of C. quinoa
showed typical symptoms 10 dpi and were densely covered with pycnidia. At 15 dpi, typical symptoms appeared on the stems of plants inoculated in outdoor conditions. Detached
inoculated leaves of C. quinoa developed visible symptoms 8 dpi and were grayish white.
However, necrotic lesions are rarely seen on the leaves in the field [137] (Figure 14A).
Figure 14. Typical symptoms of quinoa black stem in the fields of China. (A) Symptoms induced by inoculation of A. caulina
on C. quinoa (left of midrib) and on C. album (right of midrib). (B) 10 dpi diamond shaped lesion on quinoa stem with
presence of pycnidia. (C) Necrotic quinoa stem prior to lodging; (D) morphological characteristics of conidia and pycnidia
of A. caulina. Source: illustrations based on pictures from Yin et al. [114].
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Quinoa black stem primarily infects the stem; lesions are recorded at the flowering
stage up to maturity. Symptoms first appear at the lower and middle parts of the stalk,
subsequently moving upwards. They are diamond-shaped, pale or tan, and present slight
depressions, as the plants are prone to drying and consequent shrinkage. The diameter of
the lesion averages 7.9 cm.
The stem lesions turn necrotic in later stages and are accompanied by abundant small
round protrusions of black pycnidia (Figure 14B,C). In severe cases, lesions wrap around
the stem, causing lodging, foliar chlorosis, leaf abscission, and the development of “empty”
and sterile grains on the panicle [137].
Quinoa black stem is considered a newly emerging disease in Chinese regions (Jingle County, Shanxi province), where the disease was severe. The incidence was around 80%
and the yield was reduced by 45% [137,140]. The fungicides mancozeb and azoxystrobin
are shown to have a strong inhibitory effect on conidia germination, whereas tebuconazole and difenoconazole were most effective towards mycelial growth in tests performed
in vitro [137].
Sixteen European countries concentrated integrative approaches for the biological
control of the weed C. album from 1994 to 1999. The European Research Programme
(COST-816) concerted the use of a combination of A. caulina with ascaulitoxin for this
purpose [141,142]. Experiments using A. caulina as a microbial herbicide were up to 70%
successful in reducing field conditions, as it was able to kill its host in one week [143–146].
3.1.3. Cockerel Eye/Quinoa Cercorporoid Leaf Spot
Quinoa leaf spot was first reported in Ecuador (2009) and given the Spanish common
name “ojo de gallo”, or cockerel eye, because of the symptoms exhibiting a dark center
and round shape. It was then associated with Cercospora spp. [147]. The genus Cercospora
was established by Fresenius (1863) and belongs to the family Mycosphaerellaceae, class
Ascomycota. A comprehensive list of cercorporoids assembled in Poland included a species
under the name Cercospora chenopodii Fresenus, 1863, found on C. album [148].
Testen et al. (2013) amplified the ITS1-2 region of strains isolated from quinoa field
plots located in Pennsylvania, USA, and identified them as Passalora dubia (Riess) U.
Braun (GenBank EF535655). Conidia were septate, hyaline, and measured 25−98 µm
long × 5−10 µm wide—with an average of six cells per conidium (Figure 15D). Disease
symptoms of leaves were round to oval with a diameter of less than 1 cm, and were brown
to gray-black with darker brown or reddish borders (Figure 15A−C). In addition to quinoa,
P. dubia has also been isolated from C. album [77,149].
Figure 15. (A,C) depict symptoms of P. dubia on leaf tissue. (B) Comparison of “cockerel eye” and leaf spot symptoms.
(D) Conidia of P. dubia. Source: pictures (A–C) provided by Testen and (D) Testen [77].
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A pathogen identified as P. dubia has been tested as a microbial herbicide for the
biocontrol of C. album in Europe. It was shown to reduce C. album’s dry weight by 20% [143].
3.1.4. Cercospora Leaf Spot Caused by Cercospora cf. chenopodii
Cercospora leaf spot, infecting quinoa in Shanxi, China, was classified as Cercospora cf.
chenopodii based on multi-loci sequencing and phylogenetic analysis using LSU rpb2 and ITS
as target genes. The qualifier “cf” indicates a provisional identification [150], even though
most diagnostic characteristics correspond to C. chenopodii. At the early onset, the lesions were
nearly round and pale yellow to light brown. Later, the lesion became grayish brown, with a
slightly elevated surface, a yellow halo, and an average diameter of 5.4 mm. The pathogen’s
conidia were observed to be septate and hyaline to brown. They were 40.01 × 7.99 µm on
average. They contain an average of four cells per conidium (Figure 16C). Spore suspensions
made in glycerin causes disease symptoms 5 dpi, spreads quickly, and produces large
yellow lesions, which causes defoliation 10 dpi. Optimum temperatures for infections are
22–26 ◦ C, with a high relative humidity (75−80%) [151]. Based on multigene phylogeny
(LSU, rpb2, ITS, cmdA, and other genes), various Passalora species have been proposed
to be re-classified as Cercospora Fresen. P. dubia is included in this phylo-group and is
considered synonymous with Cercospora cf. chenopodii [151–153].
Figure 16. (A,B) Foliar symptoms of Cercospora leaf spot. (C) Condidia of Cercospora. Source: illustrations adapted from
Yin et al. (2019) [151].
3.1.5. Quinoa Anthracnose Caused by Colletotrichum nigrum and C. truncatum
Stem lesions have been observed on quinoa plants growing in Ames, Iowa (USA).
Symptoms are recognized as oval to linear, slightly narrow at the ends, light in color,
silvery-white to dark gray, and are slightly sunken in lesions. They contain setose acervuli.
Two isolates (CQ1, CQ2) were cultured in V8 media for the subsequent examination for
their morphological characteristics and DNA barcoding [154].
CQ1 mycelia were gray, sparse and flat. They produced abundant sclerotia and
conidia. The conidia were cylindrical, hyaline, and aseptate. The size of 50 conidia
averaged 21 × 4.3 µm. CQ2 mycelia were gray to dark and fluffy. They produced abundant
sclerotia, acervuli, and conidia. The conidia were falcate, hyaline, and aseptate. The size,
averaged from 50 conidia, was 26.8 × 2.4 µm [154]. Both isolates have been identified by
multigene sequencing, and the multiple sequence alignment of vouchered CBS isolates
generated a maximum likelihood phylogenetic tree. Based on this information, CQ1 was
identified as Colletotrichum nigrum and CQ2 as Colletotrichum truncatum. The sequences’
GenBank vouchers are: MN581860, MK675238, MF682518, and MK118057 [154].
For the completion of Koch postulates, 40-day-old quinoa plants (PI 634920) were
inoculated on stems and leaves. Two weeks later, the stems showed bleached to tan
sunken areas on wounded sites. After an extra week under humid conditions, the plants
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inoculated with C. nigrum produced acervuli (asexual stage) and sclerotia, whereas C.
truncatum produced only acervuli. Infected stems were cultured in artificial media. The
morphological characteristics of grown mycelia matched those of the initial inoculum
used on the plants. Inoculated detached leaves developed brownish, circular lesions. This
disease may cause lodging and emerge in new quinoa production areas, resulting in yield
losses [154].
3.2. Fungi Identified by Morphological Approaches
3.2.1. Brown Stalk Rot
Brown stalk rot was observed in C. quinoa growing in rotation with potatoes in the
highlands of Puno, Peru, in 1974 and 1975. The organism was isolated from diseased
stems of C. quinoa bearing pycnidia. As a practical first step for identification, the alkaline
substance 1 M NaOH was added dropwise. Its purpose was to demonstrate the presence
of substance “E” (a colorless metabolite from exigua) in malt extract agar cultures of the
fungus to distinguish it from Phoma exigua var. exigua [155]. The test gave a positive result
for P. exigua var. foveata, and comparative morphological characteristics with the causal
agent of potato gangrene were carried out [156]. As both were similar and pathogenicity
tests on potatoes were positive, the quinoa brown stalk rot’s causal agent was identified
as Phoma exigua var. foveata (Foister) Boerema. Furthermore, isolates were sent to the
Dutch Protection Service and the Commonwealth Mycological Institute (UK) for final
confirmation [157].
Symptoms were described as follows: small lesions on the higher third of the stem
progress until reaching the upper part. At this stage, pycnidia are visible, the foliage wilts,
the panicle does not form grain, and the brown stalk is prone to break (Figure 17A). The
pycnidia are globose and dark brown; their size ranges between 101−116 µm in diameter.
The ostiole is 30 µm in diameter, and the pycnidiospores are hyaline, ellipsoidal, unicellular,
and biguttulated (small drop-shaped).
Figure 17. (A) Brown stalk rot. (B) Diamond-shaped symptoms bearing pycnidia. Source: illustrations adapted from Alandia et al. [4].
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Their average size ranged between 6 × 2.2 µm in artificial media and 6.8 × 2.3 µm
when coming from infected stems. Cross-inoculations, aided (and not aided) with mechanical wounds, were performed on potato plants and tubers, tomato plants, beetroot, sugar
beets, and quinoa. Quinoa plants showed symptoms 3 dpi, potatoes and tomato plants
showed foliar blight, potato tubers got black rot, whereas beetroot and sugar beets showed
no symptoms. Overall, mechanical wounds increased the rate of infection, but pycnidia
were rarely observed. The disease developed better at 3–5 ◦ C than at 15–20 ◦ C [157].
Based on in vitro experiments, it was hypothesized [157] that the highlands of South
America are the geographic origin for the potato gangrene fungus Phoma exigua var. foveata
because it is as pathogenic to potatoes as the virulent European isolates. However, on
C. quinoa and C. album, it was more pathogenic. After inoculation, it caused a brown
discolored area of rotting tissue, 1−3 cm long on both hosts, four dpi. On older leaves of
both Chenopodium spp., concentric leaf spots of 0.5−1.0 cm in diameter were visible. The
European strain caused similar spots, but one week later [158].
3.2.2. Quinoa Diamond Black Stem/“Mancha Ojival del Allo”
Diamond black stem was observed in C. quinoa in the highlands of Puno, Peru, in
1974 and 1975. The disease is primarily present in the stem, with diamond-shaped lesions
(2−3 cm), whitish to gray in the center, with brown edges and a vitreous halo. They bear
pycnidia. At a later stage, the lesions join around the stem, causing it to collapse [133]
(Figure 17B).
3.2.3. Sclerotium in Quinoa
Stem rot affecting quinoa plants was observed at the Experimental Station of Kayr’a
(Cuzco, Peru) during 1997. The mycelium was cultured, and pathogenicity tests were
carried out on three-month-old plants of quinoa, amaranth, potato, frejol, sunflower, and
Lupinus mutabilis. All plants were infected, and quinoa was the most susceptible. Morphological comparison with Sclerotinia from potatoes allowed the morphological identification
of Sclerotinia sp., currently known as Whetzelinia sp. Inoculation with ascospores was
followed by mycelial growth after 17 days. Dark sclerotia measuring between 4−9 mm
appeared five dpi in PDA cultured at 10 ◦ C. Apothecia developed 53 dpi at 16 ◦ C and
12 days later produced ascospores. The fungi caused dry rot in the stem’s neck in quinoa,
leaves wilted, and the disease moved towards the panicle [159].
3.2.4. Damping-Off
1.
2.
3.
Sensitivity of Pythium zingiberum and P. butleri oospores: Soil inoculation of oospores
of P. zingiberum and P.butleri on soil caused damping-off of susceptible C. quinoa
seedlings after ten days of incubation at 30 ◦ C [160].
Seedling damping-off caused by Fusarium avenacearum and Pytium aphanidermatum: The fungi were isolated from infected stems of quinoa seedlings grown in
a greenhouse. Microbes were morphologically described and the cultured fungi
were inoculated on C. quinoa cv. Cochabamba. Pathogenicity tests confirmed that P.
aphanidermatum and F. avenaceum were the causal agents of the damping-off of quinoa
seedlings under greenhouse conditions. The seedling infection was significantly
higher up to the first pair of leaves, showing that quinoa is most susceptible to the
pathogens before emergence. However, the sum of post-emergence damping-off was
significantly lower than that observed in sugar beets and higher than that observed in
cabbage plants, except for F. avenacearum, which also produced marked susceptibility
at the first true leaves stage. In addition to the two pathogens, Ascochyta caulina,
Fusarium spp., and Alternaria spp were also isolated from infected tissue but could not
infect quinoa seedlings during pathogenicity tests [139].
Pathogenicity tests on seedlings infected by Rhizoctonia solani and Fusarium spp.
Rhizoctonia solani was isolated from the field in Peru. Pathogenicity tests performed
in a greenhouse showed that R. solani prevented seed germination. It also created
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4.
sunken lesions on the stems of old plants at ground level. Fusarium spp. reproduced
wilting in old plants [4,161]. Quinoa seedling damping-off (Figure 18A) was observed
during field experiments conducted at the experimental station of Nihon (Japan).
It occurred from emergence until the four-leaf stage and increased under high soil
moisture conditions. Rhizoctonia spp. (Figure 18B) and Fusarium spp. (Figure 18C)
were identified morphologically from the symptomatic lesions [162].
Pathogenicity tests on seedlings caused by Sclerotium rolfsii Sacc Sclerotium rolfsii was
isolated from diseased seedlings of C. quinoa in a field of Southern California. The
susceptibility of C. quinoa to S. rolfsii was demonstrated in vitro and under greenhouse
conditions [163].
Figure 18. (A) Quinoa seedlings affected by damping-off. (B) Rhizoctonia spp hyphae. (C) Fusarium spp. spores. (D) Healthy
quinoa seedlings growing under low soil moisture conditions. Source: illustration adapted from Isobe et al. (2019) [155].
4. Chemical Control for Oomycetes and Fungi
The control of oomycete and fungal diseases continues to rely mainly upon chemical
measures for conventional agriculture. Fungicides can be effective only on a few closely
related pathogens, in which case they are designated as narrow -spectrum fungicides,
are often systemic, and usually have a single-site activity. In contrast, broad-spectrum
fungicides can control a wide range of unrelated pathogens. In the case of oomycetes,
16 chemicals with different modes of action, translocations in the plant, types of activity,
and risks of developing resistance, are available. A summary is presented in Table 7.
However, sexual recombination, resulting in high pathogenic genetic diversity, as well as
the migration rate, including low dispersal (within a few meters), increases local epidemics
and the appearance of new pathogen genotypes in local populations. These facts will
continuously affect sensitivity to fungicides, requiring the repeated adaptation of control
strategies [105]. Several different target-site fungicides can achieve the chemical control of
fungal pathogens in a mixture or in an alternating regime on the same crop. The most recent
fungicides of this type are phenyl-pyrroles (P.P. fungicides) and dimethylation inhibitors
(DMIs). They are considered the most effective chemicals registered to control diseases
caused by Ascomycetes [164–166], depicted in blue on Table 7. The table aims to provide
a general reference. The choice of fungicide is highly dependent on the availability and
conditions of the particular fields to be treated.
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Table 7. Major fungicide groups and key active ingredients, application site, and resistance risk. Adapted from Gisi and
Zierotski (2015) [105]; Lebeda and Cohen (2021) [165]; Plimmer, (2003) [166]; and Masielo et al., (2019) [164]. Rows in blue
correspond to fungicides that are effective against Ascomycetes.
Mode of
Translocation
Fungicide Group and Key
Active Ingredients
Fully
Systemic
Phenylamides:
Metalaxyl, mefenoxam,
oxadixyl, benalaxyl,
kiralaxyl, ofurace
Resistance
Risk a
High
Foliar Seed
√
Soil
Type of
Activity
Translocation
Biochemical
in Plants
Mode of Action
Preventive, Apoplastic,
curative,
symplastic,
eradicatranslamitive
nar
√
Inhibition of rRNA
synthesis
b
Partially
Systemic
Quinone outside
inhibitors:
Azoxystrobin, fenamidone,
famox,
adone, trifloxystrobin:
kresoxin-methyl,
Pyraclostrobin
NonSystemic
b Multisites:
For example, mancozeb;
chlorothalonil, copper,
cu-oxychloride,
cu-hydroxide; folpet; thiram,
chlorothalonil
Low
√
Preventive
NonSystemic
Carboxylic acid amides:
Dimethomorph, flumorph;
iprovalicarb,
benthiavalicarb;
mandipropamid
Moderate
√
Preventive
√
Apoplastic,
Preventive, symplastic,
curative
translaminar
Inhibition of
mitochondrial
respiration at the
enzyme complex
III
Preventive
Inhibition of ATP
production
Preventive, Apoplastic,
curative
symplastic,
Inhibition of spore
germination,
retardation of
mycelia
√
Fully
Systemic
Cyanoacetamide, oximes
(cymoxanil)
Moderate
NonSystemic
Dinitroanilines (fluazinam)
Moderate
Fully
Systemic
Phosphonates (fosetyl-Al)
Moderate
√
Partially
Systemic
Quinone inside respiration
inhibitors:
Cyazofamid, amisulbrom
Medium to
hight
√
Benzamides (fluopicolide)
Mod.
√
Benzamides, carboxamides
Ethaboxam, zoxamide
Low
Fully
Systemic
Systemic
Contact
Resistance
inducer
Hymexaxol
(heteroaromatics)
b Thiadiazoles
√
√
√
√
Multi-site
inhibition
Translaminar
Apoplastic,
Preventive, symplastic,
curative
translaminar
Cell wall synthesis,
Ces3A cellulose
synthase inhibition
Delocalization of
spectrin-like
proteins
Fungal RNA and
DNA syntheses
√
√
√
translaminar
apoplastic
Preventive,
curative,
Translaminar
eradicative/
√
(Etridiazole)
Acibenzolar-S-methyl.
Preventive
Inhibition of
mitochondrial
respiration at
enzyme complex
III
Preventive,
curative
Lipid structure of
Mitochondria
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Table 7. Cont.
Mode of
Translocation
Fungicide Group and Key
Active Ingredients
Resistance
Foliar Seed
Risk a
Soil
Type of
Activity
Translocation Biochemical Mode
in Plants
of Action
b
Demethylation inhibitor
fungicides (DMIs):
Imidazoles, triazolinthiones,
triazoles
prothioconazole, prochloraz,
terbuconazole,
difenoconazole
√
√
Preventive,
curative
Sterole biosynthesis
in membranes
√
√
Preventive,
curative
Signal transduction
Preventive,
eradicaApoplastic
tive
Multi-site inhibition
Affecting the
membrane
b
PP fungicides
(Phenylpyrroles)
phenylpyrroles Fludioxonil
Fully
Systemic
Carbamates: Propamocarb,
prothiocarb
Nomenclature according to Fungicide Resistance Action Committee mode of action code list, 2014, www.frac.info (accessed on 10 June
2021). b Quinone outside inhibitors and multi-sites are broad-spectrum fungicides, including activity against fungi.
a
5. Bacteria
5.1. Bacterial Leaf Spot Caused by Pseudomonas spp.
Bacterial leaf symptoms are small irregular spots both in leaves and stems. In leaves,
they turn dark brown with concentric rings and a wet halo; in stems, they become necrotic,
causing a deep lesion and wilting [133].
5.2. Bacterial Leaf Spot Caused by Pseudomonas syringae
Bacteria were isolated from symptomatic leaves and inoculates on surface sterilized
leaves of quinoa cv. Piartal. Between three to fice dpi, leaf spots were visible (Figure 19).
The bacteria colonies were identified at the species level via morphology and molecula
tools using a Bruker Daltonik MALDI Biotyper system (Germany). The coucher for the
identified bacteria was uploades to the NCBI database as txid317 [167].
Figure 19. Symptoms of bacterial leaf spot on quinoa. Source: illustration adapted from FonsecaGuerra et al. [167].
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6. Viruses
Pathogenicity assays for the identification of viruses under greenhouse conditions
require indicator plants. These plants show distinctive and consistent reactions to virus
infections. Many plant viruses can be transmitted to indicator plants via mechanical
infection or insects. Nicotiana (tobacco) and Chenopodium are hosts for a great number of
viruses [168]. Therefore, C. quinoa could be infected with the viruses that infect host plants
that grow next to it.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
Chenopodium mosaic virus: Seedlings of C. quinoa were found to contain a highly
infectious, seed-borne virus that may remain latent. The virus was restricted to
the Chenopodiaceae and was similar to the soybean mosaic virus in morphology and
physio-chemical properties [169].
Amaranthus leaf mottle virus (ALMV): Successful infections were achieved on C.
quinoa, which exhibited chlorotic local lesions and severe systemic mosaic, leaf deformation, wilting, stunning, and finally collapse of the plants. Transmission via Aphis
gossypii was confirmed 2 to 3 weeks after the 1-day inoculation access period [170].
Arracacha virus A: AVA is common in arracacha (Arracacia xanthorrhiza) in the region
of the Peruvian Andes. AVA was not transmitted by Myzus persicae, but was transmitted by the inoculation of sap and is best propagated in C. quinoa and Nicotiana
clevelandii [171–173].
Ullucus virus C: UVC is a comovirus prevalent in Ullucus tuberosus grown at high
altitudes in the Bolivian and Peruvian Andes. It was transmitted mechanically to C.
amaranticolor and C. quinoa. It caused a systemic infection. UVC was not transmitted
by either aphid species (Aphis gossypii or Myzus persicae) or through seeds of C. quinoa.
However, it was transmitted through leaf contact between infected and healthy plants,
causing chlorosis [173].
Potato virus S (PVS): Chenopodium quinoa plants displayed symptoms of PVS infection
14 days after artificial inoculation with PVS [174,175].
Potato Andean latent virus: APLV was found to infect both C. quinoa and C. amaranticolor [176].
Cucumber mosaic virus (CMV): Partially purified extracts from leaves of Phytolacca
americana caused marked inhibition of CMV infection on C. quinoa [177].
Tobacco mosaic virus: TMV has successfully infected C. quinoa [178].
Passiflora latent virus (PLV): Chenopodium quinoa plants presenting systemic symptoms after inoculation with PLV showed high concentrations of virus particles in their
cytoplasm, mitochondria, and chloroplasts [179]
Plantago asiatica mosaic virus (PIAMV): Mechanical inoculation with infected sap of
Lilium leaves on C. quinoa yielded chlorotic or necrotic local lesions [180].
Carnation latent virus: C. quinoa is an indicator species for the carnation latent
virus [181].
Chlorotic leaf spot virus: Sap inoculation on C. quinoa resulted in a satisfactory
infection [182].
7. Conclusions and Future Directions
The growing interest in quinoa has prompted research on all aspects of this crop.
From the perspective of phytopathology, it is essential to collaborate as quinoa cultivation
has been introduced to many countries worldwide and continues to enter new regions.
Therefore, it faces different challenges in each area. The impact on final seed yield has not
been quantified for many diseases yet, as they have only been identified causing symptoms
on plant tissue, but it is essential to turn our attention to this aspect.
Determining the mycobiota in quinoa grain food is of prime importance. The presence
of seed pathogens associated with mycotoxins is concerning. These secondary metabolites
are generally produced by fungi belonging to the genera (Alternaria, Aspergillus, Fusarium
and Penicillium) [183,184]. The latter two pathogens from this list have been isolated from
quinoa plants. Thus, mycotoxin production may occur in the field or during post-harvest,
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storage, or processing [185]. Indeed, a recent comparative study [183] of mycotoxin occurrence in quinoa grains cultivated in South America, and North Europe found a large
array of mycotoxins on Northern European grain. Mycotoxins were predominantly associated with Fusarium spp. (e.g., butenolid, aurofusarin, equisetin, culmorin), Alternaria spp.
(e.g., tenuazonic acid and altersetin), Cladosporium spp. (e.g., Cladosporim), and Penicillium
spp. (e.g., ochratoxin A, flavogaucin, and mycophenolic acid). Unspecific metabolites were
also found in modest amounts. Cleaning seeds provided a considerable reduction (ca.
50%) in the content of mycotoxins, but overall the North European grains had considerably
more mycotoxins compared to South American grains even after cleaning. Weather conditions, cultivation method and post-harvest treatments could explain mycotoxins array
presence differences on grain examined. The resilience of Andean grains to the growth of
mycotoxin-producing fungi could be due to their adaptation to their natural centre of origin.
Something that drastically changes when quinoa is cultivated in other latitudes [186,187]. It
could also be argued that high saponin-containing quinoa grains may prevent the growth of
fungi [188] or serve as a fungistatic. Therefore, monitoring seed quality during post-harvest
should become a routine procedure. The implementation of this practice will highlight the
fragility of organic quinoa production in new temperate environments.
It is essential to standardize the descriptions of diseases, taking into account the
following suggestions:
-
-
-
Morphological identification paired with molecular tools for accurate descriptions of
causal agents, published in scientific journals, as well as the sharing of knowledge
within quinoa networks and conferences.
The performance of inclusive pathogenicity tests and Koch’s postulates to clarify the
type of interaction observed (e.g., pathogenic, endophytic/symbiotic, or saprophytic.).
Standardized protocols for disease propagation and assessment methods for severity
after infection.
The development of strategies for seed sanitation.
There exist several research centers located in areas where quinoa is traditionally
grown, and recently a pilot global collaborative network on quinoa (GCN-Quinoa)
(www.gcn-quinoa.org, accessed on 10 June 2021) has been established [189]. These
networks primarily share knowledge on cultivation and plant breeding. Knowledge
sharing in relation to quinoa diseases should also be considered.
More research on methodologies for the rapid, high throughput screening of quinoa
seeds and plants for the presence of economically important pathogens of quinoa is
needed. This would be useful for detecting causal agents early in disease development
and ensuring certified pathogen-free quinoa seeds. Moreover, phone apps with deep
learning models for diagnosing various plant diseases and pest attacks are becoming
interesting tools, which may be useful in the future.
Author Contributions: Writing—original draft preparation, C.C.-L.; writing—review and editing,
C.C.-L., D.B.A. and C.A.; visualization, C.C.-L.; supervision, D.B.A. and C.A.; project administration,
C.C.-L. and C.A.; funding acquisition, C.C.-L. All authors have read and agreed to the published
version of the manuscript.
Funding: This research was funded by the Plurinational State of Bolivia with the Programme of
Scholarships for Scientific Sovereignty and partly by the University of Copenhagen, Denmark.
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement: Not applicable.
Acknowledgments: We express our gratitude to Anna L. Testen and Yin et al. for sharing pictures,
to Sven-Erick Jacobsen for advice and making the seed of Titicaca, Puno and Vikinga available, and
for the illustrations, we thank Ana Sofia Patiño and Lucia Little.
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Conflicts of Interest: The authors declare no conflict of interest. The funders had no role in the design
of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or
in the decision to publish the results.
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