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Citation: Friedjung Yosef, A.;
Ghazaryan, L.; Klamann, L.;
Kaufman, K.S.; Baubin, C.; Poodiack,
B.; Ran, N.; Gabay, T.; Didi-Cohen, S.;
Bog, M.; et al. Diversity and
Differentiation of Duckweed Species
from Israel. Plants 2022,11, 3326.
https://doi.org/10.3390/
plants11233326
Academic Editors: Viktor Oláh,
Klaus-Jürgen Appenroth and
K. Sowjanya Sree
Received: 11 October 2022
Accepted: 25 November 2022
Published: 1 December 2022
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plants
Article
Diversity and Differentiation of Duckweed Species from Israel
Avital Friedjung Yosef 1,*, Lusine Ghazaryan 1, Linda Klamann 1, Katherine Sarah Kaufman 1, Capucine Baubin 1,2,
Ben Poodiack 1, Noya Ran 1, Talia Gabay 1,3, Shoshana Didi-Cohen 4, Manuela Bog 5,
Inna Khozin-Goldberg 4and Osnat Gillor 1, *
1Zuckerberg Institute for Water Research, J. Blaustein Institutes for Desert Research, Ben Gurion University,
Midreshet Ben Gurion 8499000, Israel
2Department of Ecology and Evolutionary Biology, University of Colorado Boulder, Boulder, CO 80309, USA
3Department of Life Sciences, Ben-Gurion University of the Negev, Be’er Sheva 8410501, Israel
4
French Associates Institute for Agriculture and Biotechnology of Drylands, The Jacob Blaustein Institutes for
Desert Research, Ben-Gurion University of the Negev, Midreshet Ben-Gurion 8499000, Israel
5Institute of Botany and Landscape Ecology, University of Greifswald, 17489 Greifswald, Germany
*Correspondence: avitush.f.y@gmail.com (A.F.Y.); gilloro@bgu.ac.il (O.G.)
Abstract:
Duckweeds (Lemnaceae) are tiny plants that float on aquatic surfaces and are typically
isolated from temperate and equatorial regions. Yet, duckweed diversity in Mediterranean and
arid regions has been seldom explored. To address this gap in knowledge, we surveyed duckweed
diversity in Israel, an ecological junction between Mediterranean and arid climates. We searched
for duckweeds in the north and center of Israel on the surface of streams, ponds and waterholes.
We collected and isolated 27 duckweeds and characterized their morphology, molecular barcodes
(atpF-atpH and psbK-psbI) and biochemical features (protein content and fatty acids composition).
Six species were identified—Lemna minor,L. gibba and Wolffia arrhiza dominated the duckweed
populations, and together with past sightings, are suggested to be native to Israel. The fatty acid
profiles and protein content further suggest that diverged functions have attributed to different
haplotypes among the identified species. Spirodela polyrhiza,W. globosa and L. minuta were also
identified but were rarer. S. polyrhiza was previously reported in our region, thus, its current low
abundance should be revisited. However, L. minuta and W. globosa are native to America and Far East
Asia, respectively, and are invasive in Europe. We hypothesize that they may be invasive species to
our region as well, carried by migratory birds that disperse them through their migration routes. This
study indicates that the duckweed population in Israel’s aquatic environments consists of both native
and transient species.
Keywords:
duckweed; fatty acids; DNA barcoding; diversity; biogeography; nitrogen content;
protein concentration; migration
1. Introduction
The Lemnaceae (duckweeds) family comprises the world’s smallest and fastest grow-
ing seed plants [
1
]. The duckweeds are miniscule plants that float on or below the surface of
freshwater bodies. The duckweed family was found throughout the globe, except for polar
regions, and was classified into five genera with 36 species [
2
]. Representatives of three
genera contain one or few tiny roots emerging from the fronds (Spirodela, Landoltia and
Lemna), while the two remaining genera are rootless and smaller (Wolffiella and Wolffia) [
1
].
These diminutive plants have gone through an extreme reduction in body size, with some
species less than 0.5 mm in size, thereby minimizing the need for non-photosynthetic
organs and selecting for rapid multiplication through budding [
3
]. As a consequence of
their fast growth rate, biomass production is high, providing practical applications to the
duckweeds in food [4,5], feed [6], water treatment [7,8] and biotechnology [9,10].
The first attempts to classify duckweed were based on their morphology [
1
]. However,
due to the duckweed’s diminutive size and organ reduction, morphological and anatomical
Plants 2022,11, 3326. https://doi.org/10.3390/plants11233326 https://www.mdpi.com/journal/plants
Plants 2022,11, 3326 2 of 13
classifications are challenging. Therefore, over the years, there was an attempt to classify
duckweeds based on the chemical composition of the flavonoids [
11
], isoforms of enzymes
(allozymes) [
12
] and fatty acids [
13
]. With the advancement of molecular taxonomy, molec-
ular methods of identification have been developed, including molecular fingerprinting
and sequencing [
14
]. The DNA-based molecular identification is based on the polymor-
phisms of target non-coding intron and gene spacer regions, mainly within the chloroplast
genome [
15
,
16
]. DNA markers are considered the most reliable method for species classi-
fication and have been demonstrated to be capable of detecting polymorphisms among
haplotypes of the same species [
2
]. The molecularly detected polymorphisms are sup-
ported by different physiological properties, including growth rate [
3
], protein and starch
content [
17
], metabolite abundance [
13
] and turion formation [
18
]. Therefore, haplotype
identification requires a combination of molecular and physiological-based methods [19].
Reliable identification methods are vital to establish the biogeography of duckweeds.
Early studies have reported species-dependent biodiversity, ranging from regional to global
distribution, with some species showing a broad distribution, while others were restricted
to certain regions [
16
]. Yet, a species’ global dispersal could result from the generalization of
unique haplotypes within a species. In fact, it was reported that dispersal was not directly
linked to taxonomy as closely related species—even haplotypes of the same species—were
detected oceans apart [16,20].
Duckweed growth is mostly vegetative, whereas flowering and seed generation, i.e.,
sexual reproduction, is rarer [
21
]. The survival of duckweeds during winter in temperate
regions is not only dependent on seeds; this is further evidenced by various duckweeds
reportedly sinking to the bottom of water bodies, consequently morphing into turions that
can survive freezing [
18
]. Moreover, dispersal was also attributed to biotic vectors, for
example via migrating waterfowl that can carry the small duckweeds over great distances
in their gastrointestinal tract or attached to their body [
22
,
23
]. When introduced to new
aquatic habitats, the duckweed’s fast vegetative growth facilitates their propagation and
establishment [23].
The study of duckweed diversity in the context of biogeographical distribution is
relevant to Israel because it is a meeting point between three continents: Africa, Asia and
Europe, thereby forming a transitional region between arid and Mediterranean climates [
24
].
Moreover, the Jordan valley in the east of Israel is extended from the African Rift and
serves as an important hub for migratory birds that winter in Africa and summer in
Europe [
25
]. It is a part of the Afro-Palaearctic bird migration system, the largest land bird
migration system in the world [
26
]. In spite of its small size, Israel’s location between the
Mediterranean Sea in the west and the Arabian Deserts in the east forms an ecological
corridor and a bottleneck in the birds’ flight path, making it an essential stop-over site
during migration. The bird’s stopover sites provide an opportunity for hitchhiking plants,
such as duckweeds, to establish in new environments [
27
]. Nevertheless, the diversity of
Israel’s duckweeds was never systematically investigated, though sightings of duckweed
have been documented. The following Lemnaceae species have been reported in Israel: in
the genus Lemna L. trisulca,L. gibba,L. aequinoctialis and L. minor; in the genus Spirodela:
S. polyrhiza; and in the genus Wolffia:W. arrhiza and W. globosa were reported (https://flora.
org.il/plants/systematics/lemnaceae/ (accessed on 5 September 2022)). The following
species are listed as endangered: W. arrhiza and S. polyrhiza (https://redlist.parks.org.il/
plants/list/ (accessed on 5 September 2022)). The species W. globosa was reported but
considered an invador. However, the difficulty of identifying duckweeds based solely on
their morphology, questions the reliability of these observations.
In this study we conducted a systematic survey of duckweeds in northern and central
Israel by following past sightings of duckweeds. This involved sampling the aquatic
plants, then isolating them in the lab and identifying them through morphology, molecular
methods, as well as biochemical features including fatty acid composition and nitrogen
content. We hypothesized that the duckweed diversity in these sites would reflect the
species reported in Africa, Europe and Central Asia, following birds’ migration routes.
Plants 2022,11, 3326 3 of 13
2. Materials and Methods
2.1. Survey of Duckweed Strains
During June 2021, duckweed species were collected from ponds, springs, streams and
waterholes in northern and central Israel (Galilee, Hula Valley, Golan Heights and Sharon).
The survey locations were selected based on previous observations from the last century
taken from the Israel Nature and Parks Authority database (https://redlist.parks.org.il/
plants/list/ (accessed on 11 September 2022)).
Duckweed plants were detected in 24 of the 67 reported locations detailed in the database.
The verified duckweed locations are depicted in Figure 1and detailed in Table S1.
Plants 2022, 11, x FOR PEER REVIEW 3 of 13
In this study we conducted a systematic survey of duckweeds in northern and central
Israel by following past sightings of duckweeds. This involved sampling the aquatic
plants, then isolating them in the lab and identifying them through morphology, molecu-
lar methods, as well as biochemical features including fatty acid composition and nitrogen
content. We hypothesized that the duckweed diversity in these sites would reflect the
species reported in Africa, Europe and Central Asia, following birds’ migration routes.
2. Materials and Methods
2.1. Survey of Duckweed Strains
During June 2021, duckweed species were collected from ponds, springs, streams
and waterholes in northern and central Israel (Galilee, Hula Valley, Golan Heights and
Sharon). The survey locations were selected based on previous observations from the last
century taken from the Israel Nature and Parks Authority database
(https://redlist.parks.org.il/plants/list/ (accessed on 11 September 2022)).
Duckweed plants were detected in 24 of the 67 reported locations detailed in the da-
tabase. The verified duckweed locations are depicted in Figure 1 and detailed in Table S1.
Figure 1. Confirmed duckweed sightings in northern and central Israel. White paint indicates the
areas where the survey was conducted. Scale bar is 1:20,000.
Figure 1.
Confirmed duckweed sightings in northern and central Israel. White paint indicates the
areas where the survey was conducted. Scale bar is 1:20,000.
2.2. Duckweed Collection
Plant samples were collected in duplicates in 100 mL plastic containers. The containers
were stored in a refrigerated cooler (~10
◦
C) and transported to the lab up to 48 h after
collection. For each confirmed collection site, the pH value of the water was measured by
litmus paper and the results are detailed in Table S2. In the laboratory, electric conductivity
(indicating water salinity) was measured using a conductivity meter (Cole-Parmer EW-
19820-10, Vernon Hills, IL, USA) and the results are listed in Table S2.
Plants 2022,11, 3326 4 of 13
2.3. Cultivation
In the laboratory, the collected duckweeds were sorted according to their morphol-
ogy and each isolate was sterilized by rinsing the separated fronds with a 2% sodium
hypochlorite solution (NaClO) for 2 min. Single fronds were picked and cultured in
0.5
×
Schenk and Hildebrandt (SH) basal salt mixture (Sigma–Aldrich, St. Louis, MI, USA)
supplemented by 1% sucrose at pH 5.8. The plants were grown in a controlled climate
chamber under 25
◦
C, 16 h light/8 h dark cycles, and 200–250
µ
mol m
−2
s
−1
light intensity.
Sterilization was repeated until a single unique isolate was detected. From each unique
isolate, a single sterile frond was retrieved and cultured in 0.5
×
SH agar with 0.5% sucrose
and supplemented with 100 mg L−1cefotaxime (Sigma) to avoid fungal contamination.
2.4. Morphological Identification
The morphology of the isolates was assessed according to Landolt 1986 [
1
] as well as
Les et al. [
11
]. An M205 FCA fluorescence stereo microscope (Leica, Wetzlar, Germany) and
Axio Imager 2 light microscope (Zeiss, Jena, Germany) was used.
2.5. DNA Extraction, Fragment Amplification and Sequencing
The isolates were cultivated as described above for 10–14 days then total DNA was
extracted using DNeasy Plant Pro kit (Qiagen, Hilden, Germany) following the manufac-
turer’s instructions. The extracted DNA was used as a template to amplify two plastid bar-
code loci of noncoding intergenic spacers: atpF-atpH (5
0
-ACTCGCACACACTCCCTTTCC-3
0
and 5
0
-GCTTTTATGGAAGCTTTAACAAT-3
0
) and psbK-psbI (5
0
-TTAGCATTTGTT TGGCA
AG-3
0
and 5
0
-AAGTTTGAGAGTAAGCAT-3
0
). The amplification was performed as follows:
95
◦
C pre-denatured for 3 min, followed by 35 cycles of 95
◦
C for 45 s, 55
◦
C for 45 s, 72
◦
C
for 45 s, and a further extension at 72
◦
C for 10 s. Purification of the resulting amplicons
was carried out using the AccuPrep
®
PCR/Gel Purification Kit (Bioneer, Daejeon, S. Korea),
according to the manufacturer’s instructions. The purified PCR fragments were sequenced
at McLab (San Francisco, CA, USA).
2.6. DNA Barcoding Analysis
DNA sequence alignment was generated using Geneious Prime version 2022.1.1
(http://www.geneious.com/prime/ (accessed on 15 September 2022)). Blast analysis was
performed using NCBI database (https://www.ncbi.nlm.nih.gov/ (accessed on 17 Septem-
ber 2022)) and Rutgers database (http://epigenome.rutgers.edu/cgi-bin/duckweed/blast.
cgi (accessed on 17 September 2022)). Duckweed species reference sequences of the two
loci atpF-atpH and psbK-psbI were taken from the NCBI database and added to the tree
analysis. Multiple alignments of both loci were performed using MUSCLE Alignment
(https://www.ebi.ac.uk/Tools/msa/muscle/ (accessed on 20 September 2022)). A phy-
logenetic tree was constructed using Geneious Tree Builder using the Neighbor-Joining
method with Tamura-Nei as the genetic distance model. Support values were calculated
using bootstrapping with 1000 reiterations.
2.7. Fatty Acids Analysis
Plant fatty acid composition and content were analyzed using a direct transmethylation
procedure. Plants were grown as described above for 14–21 days. After harvesting, the
cultures were placed in 20
◦
C for 12 h, then dried for 48 h in a lyophilizer (VirTis, Gardiner,
NY, USA). The dry material was ground (ULTRA-TURRAX, IKA, Merck) for 2 min at
6000 rpm. A total of ~10 mg of freeze-dried biomass was used in duplicate for analysis.
Cellular fatty acids were converted into methyl esters (FAMEs) by incubation in 2 mL
of 2% H
2
SO
4
in dry methanol (v/v) for 1.5 h at 90
◦
C with continuous stirring under
Argon gas atmosphere. Myristic acid (C
17:0
) was used as an internal standard for FAME
quantification. The reaction was terminated by the addition of 1 mL of water. A total of
1 mL of Hexane (Sigma) was then added for phase separation and extraction of FAMEs.
Hexane fractions were evaporated under N
2
gas flow and resuspended in 400
µ
L of hexane.
Plants 2022,11, 3326 5 of 13
FAMEs were analyzed by gas chromatography coupled with flame ionization detection
(GC-FID) on a TRACE Ultra Gas Chromatograph (Thermo Electron, Milan, Italy) equipped
with a programmed temperature vaporizing injector, a flame-ionization detector (FID) and
a SUPELCO WAX 10 capillary column (L
×
I.D. 30 m
×
0.25 mm, df 0.25
µ
m, Sigma), using
a temperature gradient as follows: 1 min at 130
◦
C, 8 min of linear gradient to 220
◦
C and
10 min at 220
◦
C. Helium was used as a carrier gas. Retention times of FAMEs were
compared with those of available commercial FAMEs standards (in-house library), and the
literature data [28].
2.8. Nitrogen Content Analysis
For the nitrogen content analysis, plants were grown and dried as described above. For
each sample, 2–3 mg of dry material was taken in triplicate. A Flash Smart elemental analyzer
(OEA 2000, Thermo Fisher Scientific, Waltham, MA, USA) was used for the analysis. The
nitrogen concentration results were adjusted to 1 g of dry weight. The estimation of protein was
carried out as is commonly achieved by multiplying total nitrogen by the numeric factor 6.25
but with the addition of a correction factor specific to plants [
29
]. To validate our results, we
measured the nitrogen concentrations in the same samples with FT-MIR (Fourier Transform
Midinfrared Spectroscopy) described in the Supplementary Data (File S1).
3. Results
3.1. Distribution of Duckweed Species in Israel
In total, 24 locations in the north and center of Israel were found to have duckweed
(Figure 1and Table S1). These included 14 locations in the Golan Heights, five in the
Hula Valley, two in the Galilee, and three in the Sharon. Plant samples were identified
morphologically under a microscope (Table 1). Three species of Lemna have been identi-
fied: L. gibba at highest occurrence (seven observations), L. minor (five observations) and
L. minuta (three observations). Two species of Wolffia have been identified: W. arrhiza at
high occurrence (seven observations) and W. globosa at low occurrence (one observation).
In addition, the species S. polyrhiza was observed once.
Table 1.
Identification characteristics of duckweed species found in this study [
1
,
2
]. Images were
taken using an epi-fluorescent microscope.
Species Location No of Strains Morphology General Occurrence Micrograph
Wolffia
arrhiza
Golan
Heights 9
0.5–1.5 mm long, 0.4–1.2 mm wide;
ellipsoid to spherical; upper surface
is convex, opaque, bright green,
with its greatest width slightly
below the water surface; no veins;
30–100 stomata; no roots.
Widely distributed in
temperate regions; native to
Europe, South Africa;
invasive in Brazil, Japan,
and North America.
Plants 2022, 11, x FOR PEER REVIEW 5 of 13
the cultures were placed in 20 °C for 12 h, then dried for 48 h in a lyophilizer (VirTis,
Gardiner, NY, USA). The dry material was ground (ULTRA-TURRAX, IKA, Merck) for 2
min at 6000 rpm. A total of ~10 mg of freeze-dried biomass was used in duplicate for
analysis. Cellular fatty acids were converted into methyl esters (FAMEs) by incubation in
2 mL of 2% H
2
SO
4
in dry methanol (v/v) for 1.5 h at 90 °C with continuous stirring under
Argon gas atmosphere. Myristic acid (C
17:0
) was used as an internal standard for FAME
quantification. The reaction was terminated by the addition of 1 mL of water. A total of 1
mL of Hexane (Sigma) was then added for phase separation and extraction of FAMEs.
Hexane fractions were evaporated under N
2
gas flow and resuspended in 400 µL of hex-
ane. FAMEs were analyzed by gas chromatography coupled with flame ionization detec-
tion (GC-FID) on a TRACE Ultra Gas Chromatograph (Thermo Electron, Milan, Italy)
equipped with a programmed temperature vaporizing injector, a flame-ionization detec-
tor (FID) and a SUPELCO WAX 10 capillary column (L × I.D. 30 m × 0.25 mm, df 0.25 µm,
Sigma), using a temperature gradient as follows: 1 min at 130 °C, 8 min of linear gradient
to 220 °C and 10 min at 220 °C. Helium was used as a carrier gas. Retention times of
FAMEs were compared with those of available commercial FAMEs standards (in-house
library), and the literature data [28].
2.8. Nitrogen Content Analysis
For the nitrogen content analysis, plants were grown and dried as described above.
For each sample, 2–3 mg of dry material was taken in triplicate. A Flash Smart elemental
analyzer (OEA 2000, Thermo Fisher Scientific, Waltham, MS, USA) was used for the anal-
ysis. The nitrogen concentration results were adjusted to 1 g of dry weight. The estimation
of protein was carried out as is commonly achieved by multiplying total nitrogen by the
numeric factor 6.25 but with the addition of a correction factor specific to plants [29]. To
validate our results, we measured the nitrogen concentrations in the same samples with
FT-MIR (Fourier Transform Midinfrared Spectroscopy) described in the Supplementary
Data (File S1).
3. Results
3.1. Distribution of Duckweed Species in Israel
In total, 24 locations in the north and center of Israel were found to have duckweed
(Figure 1 and Table S1). These included 14 locations in the Golan Heights, five in the Hula
Valley, two in the Galilee, and three in the Sharon. Plant samples were identified morpho-
logically under a microscope (Table 1). Three species of Lemna have been identified: L.
gibba at highest occurrence (seven observations), L. minor (five observations) and L. minuta
(three observations). Two species of Wolffia have been identified: W. arrhiza at high occur-
rence (seven observations) and W. globosa at low occurrence (one observation). In addition,
the species S. polyrhiza was observed once.
Table 1. Identification characteristics of duckweed species found in this study [1,2]. Images were
taken using an epi-fluorescent microscope.
Spe-
cies Location No of
Strains
Morphology General Occurrence Micrograph
Wolffia
arrhiza Golan Heights 9
0.5–1.5 mm long, 0.4–1.2 mm wide; ellipsoid to spherical; up-
per surface is convex, opaque, bright green, with its greatest
width slightly below the water surface; no veins; 30–100 sto-
mata; no roots.
Widely distributed in temper-
ate regions; native to Europe,
South Africa; invasive in Brazil,
J
apan, and North America.
Wolffia
g
lobosa HaSharon 2
0. 4–0.9 mm long, 0.3–0.6 mm wide; ellipsoid; upper surface
convex, translucent pale green, with its greatest width well be-
low the water surface; no veins; 8–25 stomata; no roots.
Tropical, subtropical, and
warm temperate regions; native
to eastern and southeast Asia
and Africa; invasive in North
America.
Wolffia
globosa HaSharon 2
0. 4–0.9 mm long, 0.3–0.6 mm wide;
ellipsoid; upper surface convex,
translucent pale green, with its
greatest width well below the water
surface; no veins; 8–25 stomata;
no roots.
Tropical, subtropical, and
warm temperate regions;
native to eastern and
southeast Asia and Africa;
invasive in North America.
Plants 2022, 11, x FOR PEER REVIEW 5 of 13
the cultures were placed in 20 °C for 12 h, then dried for 48 h in a lyophilizer (VirTis,
Gardiner, NY, USA). The dry material was ground (ULTRA-TURRAX, IKA, Merck) for 2
min at 6000 rpm. A total of ~10 mg of freeze-dried biomass was used in duplicate for
analysis. Cellular fatty acids were converted into methyl esters (FAMEs) by incubation in
2 mL of 2% H
2
SO
4
in dry methanol (v/v) for 1.5 h at 90 °C with continuous stirring under
Argon gas atmosphere. Myristic acid (C
17:0
) was used as an internal standard for FAME
quantification. The reaction was terminated by the addition of 1 mL of water. A total of 1
mL of Hexane (Sigma) was then added for phase separation and extraction of FAMEs.
Hexane fractions were evaporated under N
2
gas flow and resuspended in 400 µL of hex-
ane. FAMEs were analyzed by gas chromatography coupled with flame ionization detec-
tion (GC-FID) on a TRACE Ultra Gas Chromatograph (Thermo Electron, Milan, Italy)
equipped with a programmed temperature vaporizing injector, a flame-ionization detec-
tor (FID) and a SUPELCO WAX 10 capillary column (L × I.D. 30 m × 0.25 mm, df 0.25 µm,
Sigma), using a temperature gradient as follows: 1 min at 130 °C, 8 min of linear gradient
to 220 °C and 10 min at 220 °C. Helium was used as a carrier gas. Retention times of
FAMEs were compared with those of available commercial FAMEs standards (in-house
library), and the literature data [28].
2.8. Nitrogen Content Analysis
For the nitrogen content analysis, plants were grown and dried as described above.
For each sample, 2–3 mg of dry material was taken in triplicate. A Flash Smart elemental
analyzer (OEA 2000, Thermo Fisher Scientific, Waltham, MS, USA) was used for the anal-
ysis. The nitrogen concentration results were adjusted to 1 g of dry weight. The estimation
of protein was carried out as is commonly achieved by multiplying total nitrogen by the
numeric factor 6.25 but with the addition of a correction factor specific to plants [29]. To
validate our results, we measured the nitrogen concentrations in the same samples with
FT-MIR (Fourier Transform Midinfrared Spectroscopy) described in the Supplementary
Data (File S1).
3. Results
3.1. Distribution of Duckweed Species in Israel
In total, 24 locations in the north and center of Israel were found to have duckweed
(Figure 1 and Table S1). These included 14 locations in the Golan Heights, five in the Hula
Valley, two in the Galilee, and three in the Sharon. Plant samples were identified morpho-
logically under a microscope (Table 1). Three species of Lemna have been identified: L.
gibba at highest occurrence (seven observations), L. minor (five observations) and L. minuta
(three observations). Two species of Wolffia have been identified: W. arrhiza at high occur-
rence (seven observations) and W. globosa at low occurrence (one observation). In addition,
the species S. polyrhiza was observed once.
Table 1. Identification characteristics of duckweed species found in this study [1,2]. Images were
taken using an epi-fluorescent microscope.
Spe-
cies Location No of
Strains
Morphology General Occurrence Micrograph
Wolffia
arrhiza Golan Heights 9
0.5–1.5 mm long, 0.4–1.2 mm wide; ellipsoid to spherical; up-
per surface is convex, opaque, bright green, with its greatest
width slightly below the water surface; no veins; 30–100 sto-
mata; no roots.
Widely distributed in temper-
ate regions; native to Europe,
South Africa; invasive in Brazil,
J
apan, and North America.
Wolffia
g
lobosa HaSharon 2
0. 4–0.9 mm long, 0.3–0.6 mm wide; ellipsoid; upper surface
convex, translucent pale green, with its greatest width well be-
low the water surface; no veins; 8–25 stomata; no roots.
Tropical, subtropical, and
warm temperate regions; native
to eastern and southeast Asia
and Africa; invasive in North
America.
Lemna
gibba
Golan
Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower
surface of the fronds is usually
gibbous; 4–5 veins extending from
the nodes; >100 stomata; 1 root;
difficult to identify due to high
polymorphism.
Worldwide except Australia
Plants 2022, 11, x FOR PEER REVIEW 6 of 13
Lemna
g
ibba
Golan Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower surface of the fronds is
usually gibbous; 4–5 veins extending from the nodes; >100 sto-
mata; 1 root; difficult to identify due to high polymorphism.
Worldwide except Australia
Lemna
minor
Galilee, Hula Val-
ley, HaSharon 5
1–10 mm long, 6–7 mm wide; upper surface shiny green, occa-
sionally reddish; usually 3 veins, rarely 4–5; >100 stomata; 1
root.
Cooler oceanic regions; native
to North America, Europe, Af-
rica, and Western Asia.
Lemna
minuta
Golan Heights,
Hula Valley 3
0.8–4 mm long, 0.5–2.5 mm wide; forming colonies of 2–4
fronds; circular with a slightly asymmetrical base; one vein,
not very distinct; ~30 stomata; 1 root.
Temperate and subtropical re-
gions, dry to moderately humid
climate; native to America; in-
vasive in Japan and Europe
Spi-
rodela
p
oly-
rhiza
Golan Heights 1
Largest duckweed: 1.5–10 mm long, 1.5–8 mm wide; usually
thin fronds, rarely gibbous; maximum 16 veins; >100 stomata;
7–21 roots
Worldwide
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and psbK-
psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed af-
ter multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes col-
lected in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype
that was isolated from the Golan Hights, Israel [5]) and additional reference sequences (S.
polyrhiza 7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor 7123)
taken from the NCBI database (one for each species).
Plants 2022,11, 3326 6 of 13
Table 1. Cont.
Species Location No of Strains Morphology General Occurrence Micrograph
Lemna
minor
Galilee,
Hula Valley,
HaSharon
5
1–10 mm long, 6–7 mm wide; upper
surface shiny green, occasionally
reddish; usually 3 veins, rarely 4–5;
>100 stomata; 1 root.
Cooler oceanic regions;
native to North America,
Europe, Africa, and
Western Asia.
Plants 2022, 11, x FOR PEER REVIEW 6 of 13
Lemna
g
ibba
Golan Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower surface of the fronds is
usually gibbous; 4–5 veins extending from the nodes; >100 sto-
mata; 1 root; difficult to identify due to high polymorphism.
Worldwide except Australia
Lemna
minor
Galilee, Hula Val-
ley, HaSharon 5
1–10 mm long, 6–7 mm wide; upper surface shiny green, occa-
sionally reddish; usually 3 veins, rarely 4–5; >100 stomata; 1
root.
Cooler oceanic regions; native
to North America, Europe, Af-
rica, and Western Asia.
Lemna
minuta
Golan Heights,
Hula Valley 3
0.8–4 mm long, 0.5–2.5 mm wide; forming colonies of 2–4
fronds; circular with a slightly asymmetrical base; one vein,
not very distinct; ~30 stomata; 1 root.
Temperate and subtropical re-
gions, dry to moderately humid
climate; native to America; in-
vasive in Japan and Europe
Spi-
rodela
p
oly-
rhiza
Golan Heights 1
Largest duckweed: 1.5–10 mm long, 1.5–8 mm wide; usually
thin fronds, rarely gibbous; maximum 16 veins; >100 stomata;
7–21 roots
Worldwide
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and psbK-
psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed af-
ter multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes col-
lected in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype
that was isolated from the Golan Hights, Israel [5]) and additional reference sequences (S.
polyrhiza 7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor 7123)
taken from the NCBI database (one for each species).
Lemna
minuta
Golan
Heights,
Hula Valley
3
0.8–4 mm long, 0.5–2.5 mm wide;
forming colonies of 2–4 fronds;
circular with a slightly
asymmetrical base; one vein, not
very distinct; ~30 stomata; 1 root.
Temperate and subtropical
regions, dry to moderately
humid climate; native to
America; invasive in Japan
and Europe
Plants 2022, 11, x FOR PEER REVIEW 6 of 13
Lemna
g
ibba
Golan Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower surface of the fronds is
usually gibbous; 4–5 veins extending from the nodes; >100 sto-
mata; 1 root; difficult to identify due to high polymorphism.
Worldwide except Australia
Lemna
minor
Galilee, Hula Val-
ley, HaSharon 5
1–10 mm long, 6–7 mm wide; upper surface shiny green, occa-
sionally reddish; usually 3 veins, rarely 4–5; >100 stomata; 1
root.
Cooler oceanic regions; native
to North America, Europe, Af-
rica, and Western Asia.
Lemna
minuta
Golan Heights,
Hula Valley 3
0.8–4 mm long, 0.5–2.5 mm wide; forming colonies of 2–4
fronds; circular with a slightly asymmetrical base; one vein,
not very distinct; ~30 stomata; 1 root.
Temperate and subtropical re-
gions, dry to moderately humid
climate; native to America; in-
vasive in Japan and Europe
Spi-
rodela
p
oly-
rhiza
Golan Heights 1
Largest duckweed: 1.5–10 mm long, 1.5–8 mm wide; usually
thin fronds, rarely gibbous; maximum 16 veins; >100 stomata;
7–21 roots
Worldwide
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and psbK-
psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed af-
ter multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes col-
lected in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype
that was isolated from the Golan Hights, Israel [5]) and additional reference sequences (S.
polyrhiza 7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor 7123)
taken from the NCBI database (one for each species).
Spirodela
polyrhiza
Golan
Heights 1
Largest duckweed: 1.5–10 mm long,
1.5–8 mm wide; usually thin fronds,
rarely gibbous; maximum 16 veins;
>100 stomata; 7–21 roots
Worldwide
Plants 2022, 11, x FOR PEER REVIEW 6 of 13
Lemna
g
ibba
Golan Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower surface of the fronds is
usually gibbous; 4–5 veins extending from the nodes; >100 sto-
mata; 1 root; difficult to identify due to high polymorphism.
Worldwide except Australia
Lemna
minor
Galilee, Hula Val-
ley, HaSharon 5
1–10 mm long, 6–7 mm wide; upper surface shiny green, occa-
sionally reddish; usually 3 veins, rarely 4–5; >100 stomata; 1
root.
Cooler oceanic regions; native
to North America, Europe, Af-
rica, and Western Asia.
Lemna
minuta
Golan Heights,
Hula Valley 3
0.8–4 mm long, 0.5–2.5 mm wide; forming colonies of 2–4
fronds; circular with a slightly asymmetrical base; one vein,
not very distinct; ~30 stomata; 1 root.
Temperate and subtropical re-
gions, dry to moderately humid
climate; native to America; in-
vasive in Japan and Europe
Spi-
rodela
p
oly-
rhiza
Golan Heights 1
Largest duckweed: 1.5–10 mm long, 1.5–8 mm wide; usually
thin fronds, rarely gibbous; maximum 16 veins; >100 stomata;
7–21 roots
Worldwide
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and psbK-
psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed af-
ter multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes col-
lected in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype
that was isolated from the Golan Hights, Israel [5]) and additional reference sequences (S.
polyrhiza 7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor 7123)
taken from the NCBI database (one for each species).
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and
psbK-psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Table 2.
DNA identification of the duckweed isolates using atpF-atpH and psbK-psbI barcodes.
Identification was conducted using the NCBI database.
Lab
ID Species Region Coordinates
Seasonality of
the Water
Source
atpF-atpH Accession
No
Identity
(%)
psbK-psbI Accession
No
Identity
(%)
Strain
Identification
Strain
Identification
32b L. gibba Golan
Hights
33.137779,
35.725382 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
58 L. gibba HaSharon 32.333731,
34.876507 perennial RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
56 L. gibba Golan
Hights
33.09779,
35.81729 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
7L. gibba Golan
Hights
32.867917,
35.770194 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
43b L. gibba Hulla Vally 33.060001,
35.615137 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
43a L. gibba Hulla Vally 33.060002,
35.615138 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
31a L. gibba Golan
Heights
33.139682,
35.733806 perennial RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
31b L. gibba Golan
Heights
33.139682,
35.733806 perennial RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
30 L. gibba Golan
Heights
33.138450,
35.734019 seasonal RDSC 5504 KX212889.1 100 DW102 OM569589.1 100
45a L. minor Galille 32.912915,
35.569178 perennial K46 OM569601.1 100 K46 OM569540.1 100
42 L. minor Hulla Vally 33.015434,
35.629857 seasonal K46 OM569601.1 100 K46 OM569540.1 100
44 L. minor Hulla Vally 33.064187,
35.610817 seasonal K46 OM569601.1 100 K46 OM569540.1 100
57 L. minor HaSharon 32.363913,
34.958369 perennial K46 OM569601.1 100 K46 OM569540.1 100
45b L. minor Galille 32.912915,
35.569178 perennial K46 OM569601.1 100 K46 OM569540.1 100
55a L. minuta Golan
Hights
32.801403,
35.783032 seasonal 5573 MK516255.1 100 5573 MK516236.1 100
55b L. minuta Golan
Hights
32.801403,
35.783032 seasonal 5573 MK516255.0 100 5573 MK516236.2 100
Plants 2022,11, 3326 7 of 13
Table 2. Cont.
Lab
ID Species Region Coordinates
Seasonality of
the Water
Source
atpF-atpH Accession
No
Identity
(%)
psbK-psbI Accession
No
Identity
(%)
Strain
Identification
Strain
Identification
43s L. minuta Hulla Vally 33.060002,
35.615138 seasonal 5573 MK516255.1 100 5573 MK516236.3 100
19 S. polyrhiza Golan
Hights
32.969559,
35.820036 seasonal 7498 MN419335.1 100 RDSC 2014 OM569580.1 100
58 W. globosa HaSharon 32.333731,
34.876507 perennial DW2101-4 KJ630544.1 100 5514 MG812327.1 100
11b W. arrhiza Golan
Heights
32.894030,
35.775695 seasonal DW35 OM569550.1 100 DW35 OM569611.1 99.01
30a W. arrhiza Golan
Heights
33.138450,
35.734019 seasonal DW35 OM569550.1 100 DW35 OM569611.1 99.01
31b W. arrhiza Golan
Heights
33.139682,
35.733806 perennial DW35 OM569550.1 100 DW35 OM569611.1 99.01
55 W. arrhiza Golan
Heights
32.801403,
35.783032 seasonal DW35 OM569550.1 100 DW32 OM569610.1 99.29
32b W. arrhiza Golan
Heights
33.137779,
35.725382 seasonal DW35 OM569550.1 100 DW35 OM569611.1 99.31
30b W. arrhiza Golan
Heights
33.138451,
35.734018 seasonal DW35 OM569550.1 100
11a W. arrhiza Golan
Heights
32.895829,
35.776775 seasonal DW35 OM569550.1 100 DW35 OM569611.1 99.31
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed after
multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes collected
in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype that was
isolated from the Golan Hights, Israel [
5
]) and additional reference sequences (S. polyrhiza
7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor 7123) taken
from the NCBI database (one for each species).
Plants 2022, 11, x FOR PEER REVIEW 6 of 13
Lemna
g
ibba
Golan Heights,
Hula Valley,
HaSharon
9
1–8 mm long, ~3.5 mm wide; lower surface of the fronds is
usually gibbous; 4–5 veins extending from the nodes; >100 sto-
mata; 1 root; difficult to identify due to high polymorphism.
Worldwide except Australia
Lemna
minor
Galilee, Hula Val-
ley, HaSharon 5
1–10 mm long, 6–7 mm wide; upper surface shiny green, occa-
sionally reddish; usually 3 veins, rarely 4–5; >100 stomata; 1
root.
Cooler oceanic regions; native to
North America, Europe, Africa, and
Western Asia.
Lemna
minuta
Golan Heights,
Hula Valley 3
0.8–4 mm long, 0.5–2.5 mm wide; forming colonies of 2–4
fronds; circular with a slightly asymmetrical base; one vein,
not very distinct; ~30 stomata; 1 root.
Temperate and subtropical regions,
dry to moderately humid climate; na-
tive to America; invasive in Japan and
Europe
Spi-
rodela
p
oly-
rhiza
Golan Heights 1
Largest duckweed: 1.5–10 mm long, 1.5–8 mm wide; usually
thin fronds, rarely gibbous; maximum 16 veins; >100 stomata;
7–21 roots
Worldwide
3.2. DNA Barcoding Analysis
The sequences derived from the two markers psbK-psbI and atpF-atpH were used for
identifying the species listed in Table 2. Once the sequences had been cleaned, corrected
and aligned, they were blast-matched with sequences in the NCBI and Rutgers University
databases. The results for species identification, along with their accession numbers, are
presented in Table 2. All of the haplotypes were clearly identified as being of the same
species, although there was not always a match between the barcodes atpF-atpH and psbK-
psbI, as seen in the S. polyrhiza 19, W. arrhiza 56 and all L. gibba haplotypes.
Phylogenetic Tree
A phylogenetic tree based on psbK-psbI and atpF-atpH sequences was constructed af-
ter multiple alignment of the sequences (Figure 2) using a total of the 27 haplotypes col-
lected in this study, as well as W. globosa “Mankai” (a domesticated duckweed haplotype
that was isolated from the Golan Hights, Israel [5]) and additional reference sequences (S.
polyrhiza 7498, W. arrhiza DW35, W. globosa 8789, L. minuta 5573, L. gibba 5504, L. minor
7123) taken from the NCBI database (one for each species).
Figure 2.
Neighbor-Joining tree using the concatenated psbK-psbI and atpF-atpH sequences and the
Tamura-Nei distance. Based on 1000 reiterations. The numbers on the nodes represent the percentage
of bootstrap values. Horizontal bars indicate genetic distances. Reference sequences were retrieved
from the NCBI database and highlighted in bold.
Plants 2022,11, 3326 8 of 13
3.3. Fatty Acid Analysis
Total fatty acid content measured by GC-FID varied between 2.83 and 6.34% of dry weight
across the isolates. There were three major fatty acids that accounted for 80–90% of the total
fatty acids: palmitic acid (16:0), linoleic acid (LA, 18:2n6) and
α
-linolenic acid (ALA, 18:3n3).
The other fatty acids present at lower proportions are listed in Table 3and Table S3.
Table 3.
Fatty acid composition and content of duckweed plants collected in the current study. The
strain W. arrhiza 9528 was added to the analysis as a reference. An isomer of 16:1 and 18:1n7 are not
shown. The full table is shown in the Supplementary Data (Table S3).
Fatty Acid (% of TotalFatty Acids) ω3/ω6
18:3/18:2
TFA
(%DW)
Haplotype 16:0 16:1 16:2 16:3 18:0 18:1n9 18:2n6 18:3n6
GLA
18:3n3
ALA
18:4n3
SDA 20:0 22:0 24:0
L. gibba 58 20.56 ±
0.43
3.91 ±
0.42
2.2 ±
0.05
0.52 ±
0.13
0.92 ±
0.04
1.13 ±
0.13
11.1 ±
0.33
0.61 ±
0.05
52.88 ±
0.65
2.62 ±
0.08
0.37 ±
0.01
0.44 ±
0.04
1.13 ±
0.03 4.77 5.25
L. gibba 30 19.72 ±
0.46
4.88 ±
0.03
3.27 ±
0.05
0.75 ±
0.04
1.01 ±
0.16
1.07 ±
0.09
10.46 ±
0.17
0.89 ±
0.05
50.32 ±
0.36
3.81 ±
0
0.45 ±
0.01
0.39 ±
0.02
1.15 ±
0.04 4.81 4.39
L. gibba 31a 21.32 ±
0.05
4.28 ±
0.04
2.06 ±
0.01
0.17 ±
0.02
1.25 ±
0.01 0.78 ±011.27 ±
0.18 0.72 ±0 50.26 ±04.37 ±
0.03 0.53 ±00.57 ±
0.01
1.05 ±
0.01 4.46 3.63
L. gibba 31b 21.61 ±
0.17
5.46 ±
0.31
2.18 ±
0.02
0.23 ±
0.07
1.07 ±
0.04
1.11 ±
0.01
13.93 ±
0.05 0.39 ±048.98 ±
0.01
1.96 ±
0.03 0.39 ±00.41 ±
0.01
0.84 ±
0.02 3.52 3.88
L. gibba 32b 20.01 ±
0.42
4.65 ±
1.09
2.11 ±
0.03
0.35 ±
0.21
1.36 ±
0.05
0.94 ±
0.01
15.37 ±
0.09
0.8 ±
0.01
48.09 ±
0.21
3.21 ±
0.01
0.55 ±
0.02
0.51 ±
0.05
0.93 ±
0.02 3.13 4.10
L. gibba 43a 20.96 ±
0.23
3.69 ±
0.42
2.07 ±
0.06
0.65 ±
0.08
1.01 ±
0.08
1.47 ±
0.24
13.83 ±
0.02
0.61 ±
0.07
49.87 ±
0.91
2.06 ±
0.14
0.48 ±
0.02
0.51 ±
0.01
1.1 ±
0.19 3.60 4.36
L. gibba 43b 21.4 ±
0.31
4.56 ±
0.77
1.96 ±
0.03
0.43 ±
0.13
0.97 ±
0.01
1.3 ±
0.02
13.69 ±
0.07 0.47 ±049.92 ±
0.24
2.02 ±
0
0.39 ±
0.01
0.41 ±
0.02
0.76 ±
0.03 3.65 4.12
L. gibba 56 21.75 ±
2.27
5.39 ±
1.1
2.62 ±
0.02
0.23 ±
0.11
1.08 ±
0.31
1.08 ±
0.19
13.82 ±
1.37
0.52 ±
0.11
47.77 ±
3.37
2.35 ±
0.34
0.42 ±
0.14
0.45 ±
0.15
0.71 ±
0.2 3.49 4.09
L. gibba 721.49 ±
0.04
4.23 ±
0.22
2.24 ±
0.06
0.4 ±
0.03
1.24 ±
0.03
0.94 ±
0.01
15.3 ±
0.06
0.44 ±
0.05
47.78 ±
0.51
2.1 ±
0.03
0.53 ±
0.01
0.5 ±
0.01
1.4 ±
0.13 3.12 2.83
L. minor 42 22.73 ±
0.12
5.67 ±
0.37
1.62 ±
0.09
0.35 ±
0.06
1.8 ±
0.12
1.13 ±
0.02
16.01 ±
0.08
0.46 ±
0.01
45.8 ±
0.03
1.51 ±
0.14 0.55 ±0 0.31 ±01.01 ±
0.01 2.86 3.35
L. minor 44 19.85 ±
0.06
5.5 ±
0.01
2.01 ±
0.01
0.2 ±
0.01
1.28 ±
0.02 1.45 ±018.4 ±
0.1 0.5 ±046.75 ±
0.13
1.75 ±
0
0.31 ±
0.01
0.29 ±
0.01
0.82 ±
0.04 2.54 6.34
L. minor 45a 19.97 ±
0.33
2.89 ±
0.05
2.03 ±
0.02
0.75 ±
0.02
1.4 ±
0.04
1.69 ±
0.01
18.88 ±
0.05
0.6 ±
0.01
47.37 ±
0.39
1.88 ±
0.08
0.47 ±
0.01
0.36 ±
0.15
0.72 ±
0.78 2.51 3.77
L.minor 45b 22.49 ±
0.9
4.1 ±
0.79
2.54 ±
0.43
0.68 ±
0.37
1.4 ±
0.16
1.57 ±
0.03
17.64 ±
0.15
0.71 ±
0.05
42.74 ±
0.63
2.68 ±
0.21
0.45 ±
0.01
0.34 ±
0.14
1.16 ±
0.15 2.42 3.52
L. minor 57 18.28 ±
0.16
5.8 ±
0.18
2.77 ±
0.02
0.22 ±
0.04
0.77 ±
0.02
1.33 ±
0.01
16.1 ±
0.11
1.27 ±
0.01
48.36 ±
0.19
3.04 ±
0.01
0.17 ±
0.01 0.24 ±00.71 ±
0.03 3.00 4.19
L. minuta 43 18.48 ±
0.45
4.64 ±
0.03
2.43 ±
0.05
0.52 ±
0.05
1.24 ±
0.39
1.27 ±
0.02
14.07 ±
0.21
0.23 ±
0.21
54.43 ±
1.13
0.09 ±
0.02
0.25 ±
0.08
0.29 ±
0.05
1.13 ±
0.1 3.87 4.78
L. minuta 55a 19.92 ±
0.2
4.97 ±
0.52 1.62 ±00.31 ±
0.13
0.93 ±
0.03
1.08 ±
0.02
15.87 ±
0.14 0±053.35 ±
0.23
0.12 ±
0.11
0.15 ±
0.02
0.19 ±
0.03
0.79 ±
0.06 3.36 3.95
L. minuta 55b 20.07 ±
0.12
3.44 ±
0.07 1.6 ±00.66 ±
0.04
1.03 ±
0.02
1.35 ±
0.07
15.91 ±
0.07 0±053.51 ±
0.09
0.13 ±
0.18 0.25 ±0 0.15 ±01.17 ±
0.02 3.36 4.19
S. polyrhiza 19 23.15 ±
0.24
5.43 ±
0.37
6.11 ±
0.06
0.35 ±
0.04
2.35 ±
0.07
1.17 ±
0.04 5.23 ±00.04 ±
0.05
50.9 ±
0.48 0±00.58 ±
0.04
0.62 ±
0.01
2.61 ±
0.05 9.73 4.27
W. arrhiza 11a 23.22 ±
1.1
5.64 ±
0.58
3.64 ±
0.23
0.55 ±
0.03 1.62 ±02.17 ±
0.08
25.24 ±
0.32
0.28 ±
0.01
34.87 ±
0.64
0.1 ±
0.01
0.77 ±
0.01
0.64 ±
0.04
0.4 ±
0.03 1.38 4.38
W. arrhiza 11b 21.63 ±
0.26
5.36 ±
0.53
3.45 ±
0.12
0.59 ±
0.11
1.48 ±
0.07
1.87 ±
0.03
25.7 ±
0.02
0.15 ±
0.01
37.32 ±
0.56
0.1 ±
0.01
0.73 ±
0.01
0.53 ±
0.01
0.37 ±
0.08 1.45 4.55
W. arrhiza 30a 20.4 ±
0.01
5.32 ±
0.2
3.46 ±
0.02 0.48 ±01.17 ±
0.02
1.73 ±
0.01
22.75 ±
0.02 0.14 ±042.29 ±
0.14
0.08 ±
00.58 ±00.47 ±
0.01
0.31 ±
0.02 1.86 4.70
W. arrhiza 31b 24.28 ±
0.14
3.11 ±
0.22
3.19 ±
0.14
1.03 ±
0.14
1.35 ±
0.04
1.64 ±
0.04
23.78 ±
0.03
0.11 ±
0.1
38.74 ±
0.33
0.14 ±
0.02
0.69 ±
0.02 0.4 ±00.52 ±
0.03 1.63 3.18
W. arrhiza 55 21.85 ±
0.43
3.65 ±
0.25
3.33 ±
0.04
0.75 ±
0.04
1.37 ±
0.11
1.73 ±
0.04
25.7 ±
0.26
0.03 ±
0.01
38.97 ±
0.44
0.1 ±
0.01
0.6 ±
0.02
0.51 ±
0.04
0.34 ±
0.08 1.52 5.13
W. arrhiza 9528 19.05 ±
0.13
3.72 ±
0.46
2.48 ±
0.03
0.24 ±
0.02
1.81 ±
0.01
1.79 ±
0.01
24.29 ±
0.12
0.14 ±
0.01
44.41 ±
0.1
0.07 ±
0
0.86 ±
0.02
0.66 ±
0.05
0.29 ±
0.02 1.83 4.44
W. globosa 58 22.71 ±
0.05
4.04 ±
0.58
1.5 ±
0.05
0.35 ±
0.08 2.25 ±02.41 ±
0.01
25.02 ±
0.01
0.07 ±
0.39
39.51 ±
0.04
0.02 ±
0.02
0.53 ±
0.01
0.39 ±
0.02
0.19 ±
0.01 1.58 4.38
W. globosa Mankai 21.73 ±
0.16
4.39 ±
0.64
2.34 ±
0.02
0.74 ±
0.21
2.14 ±
0.02
1.9 ±
0.01
19.88 ±
0.27 0.14 ±044.64 ±
0.21
0.07 ±
0
0.41 ±
0.02
0.19 ±
0.01
0.33 ±
0.02 2.25 5.36
ALA (18:3n3) represented approximately 50% of the total fatty acids in most Lemna
isolates, whereas Wolffia species had lower concentrations of this major n3 PUFA with a
concurrent increase in 18:2n6. Accordingly, the ratio 18:3n3/18:2n6 attained higher values
in Lemna species. Spirodela differed from other duckweed species by the lowest percentage
of 18:2n6, resulting in the maximal 18:3n3/18:2n6 ratio. A variety of differences in fatty
acid composition between duckweed genera, including Lemna,Spirodela and Wolffia, have
Plants 2022,11, 3326 9 of 13
already been reported [
4
,
29
]. Another apparent difference in this study was the presence
of stearidonic acid (SDA,18:4n3), which is the product of a delta-6 (
∆
6) desaturation on
18:3n3. In this study, all L. minor and L. gibba isolates featured the presence of SDA at ~2%
to above 4% of total FA, as well as the detectable levels of another
∆
6 C18 PUFA,
γ
-linolenic
acid (GLA, 18:3n6). These data are in line with the presence of the
∆
6 desaturase gene in
Lemna [29] and Wolffia [30] species, enabling the biosynthesis of ∆6 C18 PUFA.
3.4. Nitrogen Analysis
Nitrogen content varied widely among strains, ranging from 2.2 to 5.4% of dry biomass
(Figure 3). There was no clear pattern with respect to the different duckweed species:
Nitrogen content appeared to vary depending on the species and even the strain. W.
globosa “Mankai” had the highest nitrogen content of 5.82%; it translates to a high protein
concentration of 25.52%–36.25%. The isolated W. globosa 58 showed a lower nitrogen
concentration. W. arrhiza produced high N concentrations in all five samples: 4.12–5.42%
nitrogen, translating to 18.28%–33.87% protein. The species L. minuta also produced high
results in two samples: between 4.01–4.54% nitrogen, translating to 17.64–28.38% protein.
Similar to W. globosa, nitrogen values were obtained in a wide range of values in the species
L. minor and L. gibba as well. S. polyrhiza yielded an average nitrogen content of 2.95%,
which translates into 12.98–18.44% protein (Figure 3). Because of the high variability, the
analysis was validated using FT-MIR spectroscopy that yielded similar results (Figure S1).
Plants 2022, 11, x FOR PEER REVIEW 9 of 13
L. minuta
55b 20.07 ± 0.12 3.44 ±
0.07 1.6 ± 0 0.66 ±
0.04
1.03 ±
0.02
1.35 ±
0.07
15.91 ±
0.07 0 ± 0 53.51 ±
0.09
0.13 ±
0.18 0.25 ± 0 0.15 ± 0 1.17 ±
0.02 3.36 4.19
S. polyrhiza
19 23.15 ± 0.24 5.43 ±
0.37
6.11 ±
0.06
0.35 ±
0.04
2.35 ±
0.07
1.17 ±
0.04 5.23 ± 0 0.04 ±
0.05
50.9 ±
0.48 0 ± 0 0.58 ±
0.04
0.62 ±
0.01
2.61 ±
0.05 9.73 4.27
W. arrhiza
11a 23.22 ± 1.1 5.64 ±
0.58
3.64 ±
0.23
0.55 ±
0.03 1.62 ± 0 2.17 ±
0.08
25.24 ±
0.32
0.28 ±
0.01
34.87 ±
0.64
0.1 ±
0.01
0.77 ±
0.01
0.64 ±
0.04
0.4 ±
0.03 1.38 4.38
W. arrhiza
11b 21.63 ± 0.26 5.36 ±
0.53
3.45 ±
0.12
0.59 ±
0.11
1.48 ±
0.07
1.87 ±
0.03
25.7 ±
0.02
0.15 ±
0.01
37.32 ±
0.56
0.1 ±
0.01
0.73 ±
0.01
0.53 ±
0.01
0.37 ±
0.08 1.45 4.55
W. arrhiza
30a 20.4 ± 0.01 5.32 ±
0.2
3.46 ±
0.02 0.48 ± 0 1.17 ±
0.02
1.73 ±
0.01
22.75 ±
0.02 0.14 ± 0 42.29 ±
0.14 0.08 ± 0 0.58 ± 0 0.47 ±
0.01
0.31 ±
0.02 1.86 4.70
W. arrhiza
31b 24.28 ± 0.14 3.11 ±
0.22
3.19 ±
0.14
1.03 ±
0.14
1.35 ±
0.04
1.64 ±
0.04
23.78 ±
0.03
0.11 ±
0.1
38.74 ±
0.33
0.14 ±
0.02
0.69 ±
0.02 0.4 ± 0 0.52 ±
0.03 1.63 3.18
W. arrhiza
55 21.85 ± 0.43 3.65 ±
0.25
3.33 ±
0.04
0.75 ±
0.04
1.37 ±
0.11
1.73 ±
0.04
25.7 ±
0.26
0.03 ±
0.01
38.97 ±
0.44
0.1 ±
0.01
0.6 ±
0.02
0.51 ±
0.04
0.34 ±
0.08 1.52 5.13
W. arrhiza
9528 19.05 ± 0.13 3.72 ±
0.46
2.48 ±
0.03
0.24 ±
0.02
1.81 ±
0.01
1.79 ±
0.01
24.29 ±
0.12
0.14 ±
0.01
44.41 ±
0.1 0.07 ± 0 0.86 ±
0.02
0.66 ±
0.05
0.29 ±
0.02 1.83 4.44
W. globosa
58 22.71 ± 0.05 4.04 ±
0.58
1.5 ±
0.05
0.35 ±
0.08 2.25 ± 0 2.41 ±
0.01
25.02 ±
0.01
0.07 ±
0.39
39.51 ±
0.04
0.02 ±
0.02
0.53 ±
0.01
0.39 ±
0.02
0.19 ±
0.01 1.58 4.38
W. globosa
Mankai 21.73 ± 0.16 4.39 ±
0.64
2.34 ±
0.02
0.74 ±
0.21
2.14 ±
0.02
1.9 ±
0.01
19.88 ±
0.27 0.14 ± 0 44.64 ±
0.21 0.07 ± 0 0.41 ±
0.02
0.19 ±
0.01
0.33 ±
0.02 2.25 5.36
3.4. Nitrogen Analysis
Nitrogen content varied widely among strains, ranging from 2.2 to 5.4% of dry bio-
mass (Figure 3). There was no clear pattern with respect to the different duckweed species:
Nitrogen content appeared to vary depending on the species and even the strain. W. glo-
bosa “Mankai” had the highest nitrogen content of 5.82%; it translates to a high protein
concentration of 25.52%–36.25%. The isolated W. globosa 58 showed a lower nitrogen con-
centration. W. arrhiza produced high N concentrations in all five samples: 4.12–5.42% ni-
trogen, translating to 18.28%–33.87% protein. The species L. minuta also produced high
results in two samples: between 4.01–4.54% nitrogen, translating to 17.64–28.38% protein.
Similar to W. globosa, nitrogen values were obtained in a wide range of values in the spe-
cies L. minor and L. gibba as well. S. polyrhiza yielded an average nitrogen content of 2.95%,
which translates into 12.98–18.44% protein (Figure 3). Because of the high variability, the
analysis was validated using FT-MIR spectroscopy that yielded similar results (Figure S1).
Figure 3. Nitrogen concentration (A) and estimated protein concentration (B) adjusted to duckweed
dry biomass, which is calculated by multiplying nitrogen concentration by 4.4–6.25. The strain W.
arrhiza 9528 was added to the analysis as a reference.
Figure 3.
Nitrogen concentration (
A
) and estimated protein concentration (
B
) adjusted to duckweed
dry biomass, which is calculated by multiplying nitrogen concentration by 4.4–6.25. The strain
W. arrhiza 9528 was added to the analysis as a reference.
4. Discussion
In a campaign to explore the diversity of duckweed in Israel, we have followed
almost 100 years of reports since 1926 (https://redlist.parks.org.il/plants/list/ (accessed on
22 September 2022)). We found duckweeds in ~40% of the seasonal and perennial water
bodies explored (confirming 24 of 67 sights). In the confirmed sites, we collected the species,
then we transferred them to the lab, there isolating and identifying the duckweeds. The
identification was based on combined approaches that included morphology, molecular
analyses and the biochemical features of the isolates (Tables 1and 3). As a result, six
species were confirmed (Figures 1and 2), with five of the identified duckweed species
previously reported in Israel. One of the identified species, L. minuta, was collected in
Israel for the first time. In this approach, the two barcode regions previously proposed [
31
],
Plants 2022,11, 3326 10 of 13
atpF-atpH and psbK-psbI, have proven to well complement the morphological identification.
Interestingly, this geographically limited study shows a pattern that was reported in
geographically broader studies. Here, the two identified Wolffia species show stronger
intraspecific differences between haplotypes than the other species [
31
], as was shown for
the W. globosa haplotypes [
14
] identified by the two chloroplast markers in Wolffia. However,
we note that the chloroplast markers do not allow for the identification of hybrids, which
may well occur between the two species of L. minor and L. gibba, as recently described [
32
].
Three of the isolated species dominated the community: L. gibba,L. minor and W. arrhiza
(accounting for 31, 17 and 31% of the isolates, respectively) and were previously sighted
in Israel (https://biogis.huji.ac.il/heb/home.html (accessed on 25 September 2022)). We
suggest that these species form the stable and established population of duckweeds in
Israel. Their continuous presence in perennial and seasonal aquatic sites across Israel
over the last century supports our hypothesis. Moreover, populations of L. gibba,L. minor
and W. arrhiza were previously reported in North Africa, the Middle East and Europe
(https://europlusmed.org/ (accessed on 25 September 2022)).
Three of the remaining species identified in this study, L. minuta,W. globosa and
S. polyrhiza, were detected at a lower abundance (Figure 1and Table 1). S. polyrhiza is
native to the Middle East and Europe [
33
] and was previously sighted in Israel (https:
//biogis.huji.ac.il/heb/home.html (accessed on 26 September 2022)). Yet, S. polyrhiza’s
low abundance detected in our survey (Figure 1and Table 1) may reflect the species’
growth inhibition by anthropogenic contamination, as was previously described [
34
,
35
].
The other two species identified in Israel, L. minuta and W. globosa, are not native to our
region and are considered invasive species in Europe (https://europlusmed.org/ (accessed
on 28 September 2022), [
36
]). L. minuta is native to America and is also an invasive
species to Europe where it was first sighted in the 1990
0
s, probably through an accidental
introduction [
36
]. L. minuta’s fast vegetative reproduction, high fitness and aggressive
competition damages European lake ecosystems by inhibiting local duckweed populations
such as L. minor [
36
]. L. minuta was spotted in the Hula Valley, a popular overnight break to
many migrating waterfowl [
25
], hence, we hypothesize that the species was inadvertently
carried by the migrating birds on their way from Europe to Africa and established there.
The species W. globosa is native to the Far East (Thailand, Cambodia and Laos [
1
]),
China [
37
], and the Indian subcontinent [
3
]). Like L. minor, it was inadvertently introduced
to Europe. Although W. globosa is the fastest propagating Angiosperm, with a doubling time
as low as 72 h [
3
], it does not aggressively compete with native species but instead resides
alongside them in water bodies across Europe [
38
]. In Israel, W. globosa was occasionally
sited [
5
] but its presence in water bodies was transient (we were unable to find the species
in the reported sites). The transient presence of W. globosa in Israel may suggest that it
cannot survive the long Mediterranean summer drought and is reintroduced to various
water bodies by birds migrating from Europe.
The molecular phylogeny of the duckweed species converged some isolates, suggest-
ing a common haplotype (Figure 2), yet the biochemical features of species, including fatty
acid profiles and nitrogen content, suggested diversification among the haplotypes (Table 3
and Figure 3). The nitrogen concentration results were validated by two independent
methods (Elemental Analyzer (Figure 3) and FT-MIR (Figure S1)). The nitrogen contents of
the haplotypes were diverse, but not always taxonomically associated. Diverging nitrogen
concentrations among duckweeds were likewise reported for various haplotypes of the
genera Wolffia [
4
,
30
] and Lemna [
39
]. There is a possibility that even if laboratory growth
conditions are controlled, the optimal growth conditions for each haplotype and species
can differ, which may have an effect on the biochemical composition, and specifically, on
the nitrogen and protein content [
4
]. Fatty acid composition was consistent with previous
duckweed surveys [
15
], showing the predominance of ALA (18:3n3) and the presence of
a considerable amount of SDA (18:4n3) mostly in Lemna species. Some Wolffia species,
yet not the ones isolated here, were shown to contain ALA and SDA (W. australiana and
W. microscopica [
30
]). The presence of SDA, whose biosynthesis requires
∆
6-desaturation, is
Plants 2022,11, 3326 11 of 13
restricted to only a few terrestrial plant families. However, this n-3 PUFA (SDA) widely
occurs in cyanobacteria and algae, as well in some duckweeds [
4
,
13
,
29
], indicating the
evolutionary radiation of the
∆
6-desaturation, and this fatty acid’s possible importance to
duckweeds. Concomitant to the diverse nitrogen contents, some haplotypes of the same
species showed variability in their fatty acid content, for instance SDA content in L. gibba
(Table 3).
Our study suggests that both stable and transient duckweed communities are present
side-by-side in Israel’s water bodies. Almost 100 years of duckweed sightings suggest
that L. gibba,L. minor and W. arrhiza inhabit the perennial water bodies but also survive
the summer drought of seasonal ponds. Resilience of duckweed under drought gained
little attention in contrast to survival during water freezing [
40
,
41
] and should be further
explored, especially considering the currently changing climate. Three additional species
were identified: S. polyrhiza,L. minuta and W. globosa. While the first was sighted in this
region, the two latter are native to America and the Far East, respectively, and considered
invasive species in Europe. We hypothesize that these species are transient in Israel, carried
by migrating waterfowl on their way from Europe to Africa and established in water bodies
as was previously proposed [
16
]. However, considering the aggressive nature of some
invading species (like L. minuta [
36
]), their potential to endanger the fragile community
of the endogenous duckweeds in Israel should be considered. However, to validate the
community composition of duckweeds in Israel, additional surveys should be conducted
covering wider spatial and temporal scales. These surveys should be accompanied by
careful isolation of the species followed by a combination of methods to identify both their
taxonomy and traits.
5. Conclusions
Here, we described the first survey of duckweed ever performed in Israel—a known
junction between Mediterranean and arid environments. We isolated 27 duckweed hap-
lotypes and used morphological and molecular approaches to identify them, resulting in
six confirmed species. However, independent of the taxonomy, the haplotypes differed in
their fatty acid profiles and protein contents. Three of the species were abundant among
sites and confirmed by past sightings, thus, were proposed as being native to Israel. The
other three species were rarer with two suspected invaders to our region. Thus, future
surveys should be conducted to establish the identity and traits of the native duckweed
communities in Mediterranean and arid regions.
Supplementary Materials:
The following supporting information can be downloaded at: https:
//www.mdpi.com/article/10.3390/plants11233326/s1, Table S1. Duckweed plants collected in
the present study—detailed location; Table S2. Water samples analysis: estimated pH and electric
conductivity (EC); Table S3. Full table of fatty acid composition and content of duckweed plants
isolates collected in the current study. The strain W. arrhiza 9528 was added to the analysis as
a reference data continuance; Figure S1. Nitrogen concentration measured by N content FT-MIT;
File S1. FTIR spectra collection.
Author Contributions:
Conceptualization, A.F.Y. and O.G.; Data collection A.F.Y., L.G., L.K., K.S.K.,
C.B., B.P., N.R. and T.G.; methodology, A.F.Y., L.G., L.K. and M.B.; investigation, A.F.Y., S.D.-C. and
I.K.-G.; resources, O.G. and I.K.-G.; data curation, A.F.Y.; writing—original draft preparation, A.F.Y.,
O.G. and I.K.-G.; writing—review and editing, A.F.Y., L.G., L.K., K.S.K., C.B., B.P., N.R., T.G., S.D.-C.,
M.B., I.K.-G. and O.G.; visualization, A.F.Y., B.P. and M.B.; supervision, O.G.; funding acquisition
O.G. and I.K.-G. All authors have read and agreed to the published version of the manuscript.
Funding: This research was funded by Israel Ministry of Agriculture grant number 16-38-0038.
Institutional Review Board Statement: Not Applicable.
Informed Consent Statement: Not Applicable.
Data Availability Statement: Not Applicable.
Plants 2022,11, 3326 12 of 13
Acknowledgments:
This research was supported by the Israel Ministry of Agriculture Grants
No 16-38-0038 to O.G. We would like to acknowledge the Negev Scholarship for the generous
support to A.F.Y. We are grateful to Appenroth for his advice and for providing duckweed strains.
We greatly appreciate Hernandez-Allica, Barrero-Sicilla, Mashanova, Espinosa-Montiel and Blachez
for the FT-MIR analyses of the nitrogen concentration. We also greatly acknowledge the help and
support from Hinoman, L.T.D. and especially Lapidot. We are grateful for the assistance given by
the Israel Nature and Park Authority and for the information they provided (Data—Scientific Data
Department, Israel Nature and Parks Authority).
Conflicts of Interest: The authors declare no conflict of interest.
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