Phytohormone profiling in an evolutionary framework

Abstract

The genomes of charophyte green algae, close relatives of land plants, typically do not show signs of developmental regulation by phytohormones. However, scattered reports of endogenous phytohormone production in these organisms exist. We performed a comprehensive analysis of multiple phytohormones in Viridiplantae, focusing mainly on charophytes. We show that auxin, salicylic acid, ethylene and tRNA-derived cytokinins including cis-zeatin are found ubiquitously in Viridiplantae. By contrast, land plants but not green algae contain the trans-zeatin type cytokinins as well as auxin and cytokinin conjugates. Charophytes occasionally produce jasmonates and abscisic acid, whereas the latter is detected consistently in land plants. Several phytohormones are excreted into the culture medium, including auxin by charophytes and cytokinins and salicylic acid by Viridiplantae in general. We note that the conservation of phytohormone biosynthesis and signaling pathways known from angiosperms does not match the capacity for phytohormone biosynthesis in Viridiplantae. Our phylogenetically guided analysis of established algal cultures provides an important insight into phytohormone biosynthesis and metabolism across Streptophyta.

Full text

Article https://doi.org/10.1038/s41467-024-47753-z
Phytohormone profiling in an evolutionary
framework
Vojtěch Schmidt
1,2,4
,RomanSkokan
1,4
, Thomas Depaepe
3
,
Katarina Kurtović
2
,SamuelHaluška
1,2
, Stanislav Vosolsobě
2
,
Roberta Vaculíková
1
, Anthony Pil
3
, Petre Ivanov Dobrev
1
, Václav Motyka
1
,
Dominique Van Der Straeten
3
&JanPetrášek
1,2
The genomes of charophyte green algae, close relatives of land plants, typi-
cally do not show signs of developmentalregulationbyphytohormones.
However, scattered reports of endogenous phytohormone production in
these organisms exist. We performed a comprehensive analysis of multiple
phytohormones in Viridiplantae, focusing mainly on charophytes. We show
that auxin, salicylic acid, ethylene and tRNA-derived cytokinins including cis-
zeatin are found ubiquitously in Viridiplantae. By contrast, land plants but not
green algae contain the trans-zeatin type cytokinins as well as auxin and
cytokinin conjugates. Charophytes occasionally produce jasmonates and
abscisic acid, whereas the latter is detected consistently in land plants. Several
phytohormones are excreted into the culture medium, including auxin by
charophytes and cytokinins and salicylic acid by Viridiplantae in general. We
note that the conservation of phytohormone biosynthesis and signaling
pathways known from angiosperms does not match the capacity for phyto-
hormone biosynthesis in Viridiplantae. Our phylogenetically guided analysis of
established algal cultures provides an important insight into phytohormone
biosynthesis and metabolism across Streptophyta.
Land plants evolved from within a group of freshwater green algae
called charophytes. Together they form the lineage Streptophyta,
which is embedded in Viridiplantae alongside the earlier-diverging
chlorophyte and prasinodermatophyte algae1. The extant charophyte
lineages are highly morphologically divergent, ranging from uni-
cellular and filamentous to multicellular with differentiated cell types2.
Phytohormones comprise several classes of endogenous,
organic substances that influence physiological processes at lower
concentrations than vitamins or nutrients3. These phytohormone
classes include auxins, cytokinins (CK), jasmonates, ethylene, absci-
sic acid (ABA), salicylic acid (SA), strigolactones (SL), gibberellins
(GA) and brassinosteroids (BR). Genomic evidence suggests that the
machinery underlying ethylene and CK signaling evolved prior to the
emergence of land plants4, but it is currently unknown whether
ethylene or CK serves as conserved signals between green algae and
land plants. The canonical perception mechanisms of auxin, ABA,
jasmonates, and SA were imposed on pre-existing transcriptional
networks in the last common ancestor of land plants4–6. Finally, the
receptors of SL, GA, and BR emerged at different points in the evo-
lution of vascular plants7–9. While the gene families underlying CK
biosynthesis predate plant terrestrialization, these are not easily
distinguishable in green algae and bryophytes from the ancient
mechanisms of tRNA modification and purine metabolism10–12. The
canonical biosynthetic pathways (as known from angiosperms) of the
other above-listed phytohormones were assembled in the last com-
mon ancestor of land plants or later in the lineage, even though
Received: 1 September 2023
Accepted: 9 April 2024
Check for updates
1
Institute of Experimental Botany of the Czech Academy of Sciences, Rozvojová 263, 165 02 Prague 6, Czechia.
2
Department of Experimental Plant Biology,
Charles University, Viničná 5, 128 44 Prague 2, Czechia.
3
Laboratory of Functional Plant Biology, Ghent University, K.L. Ledeganckstraat 35, B-9000
Ghent, Belgium.
4
These authors contributed equally: Vojtěch Schmidt, Roman Skokan. e-mail: skokan@ueb.cas.cz;petrasek@ueb.cas.cz
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homologs of certain genes are present in both charophytes and
chlorophytes5,6,13–15.
Genomic inquiries into the origins of phytohormone biosynthesis
have been paralleled by profiling of endogenous phytohormones.
First, it should be noted that the biosynthesis of many compounds
recognized as phytohormones is not restricted to Viridiplantae16–18.In
green algae, research to date has prioritized unicellular chlorophytes.
Despite their distant relationship with land plants, chlorophytes were
reported to produce nearly all the compounds known as bioactive
phytohormones19. Charophytes, the closer living relatives of land
plants, have attracted considerably less attention in phytohormone
profiling. Results obtained from a few charophyte genera collectively
attest that they can likewise produce compounds known as
phytohormones15,18,20–29. However, phytohormone profiling in green
algae and bryophytes is not without controversy, as individual studies
do not always agree on the detection of specific compounds in
equivalent biological material. Examples include abscisic acid in the
chlorophyte Draparnaldia spp.22,30, and jasmonic acid in the char-
ophyte Chara spp.26,28 and the bryophyte Marchantia
polymorpha20,21,31,32. Three studies in the charophyte Klebsormidium
spp. differed in both detection and endogenous concentrations of CK
but closely agreed on the auxin indole-3-acetic acid (IAA)18,25,27.Indivi-
dual studies tend to be limited in taxon sampling or the selection of
analyzed compounds. Moreover, multiple lineages of charophytes
have never been explored by phytohormone profiling, despite their
important phylogenetic position in plant evolutionary research33,34.A
systematic, analytical investigation of phytohormone biosynthesis and
metabolism in Streptophyta that predate transition to land has never
been performed.
To address the inconsistencies and gaps in analytical evidence,
we provide a broad screen of compounds corresponding to multiple
classes of phytohormones in a wide range of Viridiplantae. The
selected taxa represent all families of charophyte algae, but also
include several chlorophyte algae and land plants as outgroups. All
investigated organisms were cultured under closely comparable
conditions and subjected to the same analytical methods. We found
that compounds known as auxin, CK, ABA, jasmonates, SA, and
ethylene are produced in both charophytes and land plants, with
some notable distinctions between the two lineages. In addition,
several phytohormones were found to be released into the culture
media. We discuss our data in the context of current hormonomics
and genomics.
Results
Analytical design in an evolutionary framework
The algal and land plant strains were selected based on their phylo-
genetic position (Fig. 1), establishment in research as model organ-
isms, availability in axenic cultures, and ease of cultivation (see
Supplementary Data 1). Both biomass and the corresponding culture
media have been sampled at a proliferative stage of growth, but mul-
tiple selected algae were likewise sampled at the stationary growth
phase, particularly the charophytes. A liquid chromatography/mass
spectrometry (LC/MS)-based method designed to detect the broadest
possible spectrum of phytohormone compounds was applied to ana-
lyze the samples (full list of analytes in Supplementary Data 2). Addi-
tionally, the extracellular release of the gaseous phytohormone
ethylene by living algal and plant biomass was investigated using laser-
based photoacoustic spectroscopy. The results obtained were related
to the same analysis performed on blank media containing no biolo-
gical material.
Phytohormones in biomass
The profiles of phytohormones and their metabolites measured in
algal and land plant samples are summarized in Fig. 1(see Supple-
mentary Figs. 1–3 for more detailed versions). The raw dataset is
provided in Supplementary Data 3 for all compounds except ethylene,
which is detailed in Supplementary Data 4.
ABA was detected rarely in green algae, at low concentrations (pM
to units of nM), and universally in the charophyte samples that were
investigated at the stationary growth stage. By contrast, ABA was
detected consistently in the thalli of the investigated land plants (Fig. 1,
Supplementary Fig. 2). The concentrations in the lycophyte Selaginella
uncinata were similar (order of 101nM)whilethoseinbryophyteswere
lower (units of nM) compared to those previously measured in these
lineages (Supplementary Data 5). Only the lycophyte contained the
ABA catabolites phaseic and dihydrophaseic acids (PA & DPA). These
results summarily show that ABA can be produced in charophytes at
low concentrations in stationary-phase cultures, whereas it can be
regularly detected in the growing cultures of land plants. Moreover,
only the tested lycophyte showed evidence of active ABA metabolism
under our conditions.
The auxin profile in our dataset was characterized by the omni-
presence of free IAA, its catabolite 2-oxo-IAA (oxIAA), and the frequent
presence of the IAA precursor indole-3-acetamide (IAM). Land plants
differed from green algae by additionally containing the amide- and
ester-conjugates of IAA (Fig. 1), namely IAA-glutamate (IAA-Glu) and
IAA-glucosyl ester (IAA-GE). The endogenous concentrations of IAA
ranged toward the lower end compared to the available literature,
where these span the entire nanomolar scale (Supplementary Data 5).
The phenolic compound phenylacetic acid (PAA) is known to have a
weak auxin activity and is widespread in the tree of life35;itwas
accordingly omnipresent in our dataset. To conclude, we found that
free IAA, PAA, and some compounds related to both were present in all
the organisms investigated. However, only land plants showed evi-
dence of active IAA metabolism by conjugation into amides and
glucosides.
Among CK, we found that both chlorophyte and charophyte
green algae contained only the tRNA-related CK types, namely N6-(Δ2-
isopentenyl)-adenine (iP), cis-zeatin (cZ) and methylthio CK (MeS-
zeatin and MeS-iP). Land plants likewise contained these, but addi-
tionally produced the trans-zeatin (tZ) type CK and cZ/tZ-O-glucosides.
The last CK type analyzed, dihydrozeatin (DZ), as well as CK-N-gluco-
sides were absent in all strains analyzed (Fig. 1). The measured CK
concentrations varied by strain but were mostly similar between green
algae and land plants and within the ranges reported previously
(Supplementary Data 5), i.e., from pM to units of nM. In summary, we
found that all the tested Viridiplantae contained CK derived from the
post-transcriptional modifications of tRNA. However, only land plants
produced the tZ-type CK and CK-O-glucosides, the latter indicating
active CK metabolism.
The analysis of oxylipins/jasmonates (henceforth ‘jasmonates’)in
the selected Viridiplantae revealed a particularly patchy profile. A few
individual strains produced copious amounts, whereas other strains
(including all four tested chlorophytes) contained none of the tested
jasmonates (Fig. 1). The most frequently detected and simultaneously
the most abundantly produced jasmonate was dinor-12-oxo-
phytodienoic acid (dnOPDA), with two strains of Coleochaete scutata
containing particularly high (µM) endogenous concentrations (Sup-
plementary Data 3). dnOPDA was altogether found in four char-
ophytes, while its detection in Marchantia polymorpha represented
the only case of jasmonate occurrence in our tested land plants. The
charophytes Coleochaete scutata and the two strains of Klebsormi-
diophyceae constituted the only organisms thatproducedjasmonates
other than dnOPDA, namely OPDA and jasmonic acid (JA). In Kleb-
sormidiophyceae, JA was only found in the stationary-stage cultures.
Jasmonoyl-isoleucine (JA-Ile), the bioactive jasmonate of euphyllo-
phyte vascular plants15, was absent in our entire dataset. The measured
concentrations ranged from nM (JA, OPDA) to µM(dnOPDAinsome
strains). The levels of dnOPDA in Marchantia polymorpha and JA &
dnOPDA in Klebsormidium nitens in our analysis were similar to those
Article https://doi.org/10.1038/s41467-024-47753-z
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ABA
IAA
IAM
oxIAA
IAA−Glu
IAA−GE
tZ types
cZ types
Z(R)OGs
iP types
JA
OPDA
SA
PA A
BA
ABA AUXINS CYTOKININS JAs PHENOLICS
†) wild-collected (no blank)
detected at stationary phase
Marchantia
polymorpha
Spirogyra
sp.
Mougeotia
scalaris
Closterium
p-s-l complex
Mesotaenium
endlicherianum
Coleochaete
scutata
†Nitella
sp.
Chara
braunii
Klebsormidium
nitens
Interfilum
paradoxum
Chlorokybus
melkonianii
Mesostigma
viride
Desmodesmus
communis
Chlorella
sorokiniana
Trentepohlia
annulata
Picocystis
salinarum
Physcomitrium
patens
protonema
gametophores
Selaginella
uncinata
rhizophores
shoots
MeS types
oxIAA−GE
PA+DPA
dnOPDA
ETHYLENE
−4 04
biomass-to-blank ratio
log(biomass/blank)
absent in
blank
−2 2
endogenous
concentration
[pmol/gFW] < 1 1
10
100
1 000
10 000
n.d
KEY:
ethylene
emanation
[pmol/h/gFW]
&
Fig. 1 | Endogenous phytohormone compounds detected in the biomass of
green algae and land plants and ethylene emanation. Circle size denotes mean
concentration in biomass (pmol per gram fresh weight) and mean ethylene ema-
nation (rightmost column; pmol/h/gFW). No circle, compound not detected (n.d.).
Color codedenotes the ratio betweenthe concentrations measured in biomass and
blank medium (the latter containing no biological material), expressed in loga-
rithmic scale. Blue shading, compound(s) prevalent in blank. Red shading, com-
pound(s) prevalent in biomass. Black, compound(s) absent in blank. Symbols: star,
compound only detected in stationary-phase cultures; cross, wild-collected biolo-
gical material (no blank available). ABA abscisic acid, PA phaseic acid, DPA
dihydrophaseic acid, IAA indole-3-acetic acid, IAM indole-3-acetamide, oxIAA 2-
oxo-IAA, IAA-Glu IAA-glutamate, IAA-GE IAA-glucose ester, tZ trans-zeatin, cZ cis-
zeatin, Z(R)OGs zeatin (riboside)-O-glucosides; both cis-andtrans- isomers, iP N6-
(Δ2-isopentenyl)-adenine, MeS methylthio, JA jasmonic acid, OPDA 12-oxo-
phytodienoic acid, dnOPDA dinor-OPDA, SA salicylic acid, PAA phenylacetic acid,
BA benzoic acid. Minimum n= 3 for biomass (independent cultures; exact sample
size for each strain is listed in Supplementary Data 3), n=2 for blank media. Ethy-
lene measurements for both biomass and blank media were performed in n=5.
Data variation is shown in Supplementary Figs. 3 and 4 and listed in Supplemen-
tary Data 8.
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 3
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reported15, although our two studies differed in OPDA detection. In
conclusion, we observed several cases of jasmonate production in the
tested streptophytes, plausibly indicative of a conserved reaction to
the culture conditions but with lineage-specific sensitivity. dnOPDA
was detected in five of the investigated strains, while only three of
these strains additionally contained ODPA and/or JA.
SA was detected inall tested species, with no apparent differences
between land plants and green algae (Fig. 1). We likewise observed the
omnipresence of benzoic acid (BA), which can serve as an immediate
SA precursor5, but also is a more general primary metabolite36.The
measured SA concentrations ranged on the nanomolar scale, as pre-
viously reported in green algae, bryophytes, and many vascular plants
(Supplementary Data 5)5. In short, we observed that all tested Vir-
idiplantae produced SA and BA.
As ethylene is a gaseous, diffusable phytohormone, we mea-
sured its accumulation in enclosed vials containing biomass and
culture medium over a period of 6 hours using laser-based photo-
acoustic spectroscopy. A reduced sample set was analyzed com-
pared to the LC/MS screen. Both land plants and charophytes
released ethylene into the environment (Fig. 1and Supplementary
Fig. 4), although the latter at much lower amounts (1–2 orders of
magnitude). The exceptions among algae are, peculiarly, the
chlorophyte Chlorella sorokiniana and the early-diverging char-
ophyte Mesostigma viride, both of which released ethylene at levels
similar to land plants. The rate of ethylene release by Spirogyra sp.
was closely comparable to an analogous measurement performed
previously in Spirogyra pratensis29. Hence, we found that repre-
sentatives across Viridiplantae can produce ethylene. However,
land plants produced much higher amounts compared to the more
closely related charophyte lineages.
Last but not least, we note that our LC/MS method allows for the
detection of certain GA, BR, and SL (Supplementary Data 2), although
these compounds are typically measured using optimized purification,
extraction, and analysis procedures. We detected none of the analyzed
compounds under our approach, although other studies had pre-
viously reported their detection in similar biological material. This
includes GA in unicellular chlorophytes, Chara spp. and Selaginella
moellendorffii37–39, BR in unicellular chlorophytes40 and SL in
bryophytes24,41.
Phytohormones in culture and control media
Phytohormones are recognized for developmental control in the
three-dimensional bodies of land plants with differentiated tissues and
cell types. Most green algae do not possess this morphological com-
plexity, leading to discussions about whether an ancestral function of
phytohormones, if present in ancient or extant green algae, might be
facilitated via their release into the environment42,43. This motivated us
to analyze the content of phytohormone compounds not only in bio-
mass, but also in the corresponding culture media. The results are
summarized in Fig. 2a (raw data in Supplementary Data 3) and hen-
ceforth described.
CK and phenolics (SA, PAA, BA) were detected in the culture
media from the majority of investigated strains. CK levels in culture
media were lower compared to biomass. Phenolics varied in their
prevalence between culture media or biomass by the specificanalyte
and individual organism. Auxins were detected less frequently in the
culture media compared to biomass, and the ratio between the two
again varied by analyte and species. While free IAA was detected in the
culture medium of half the investigated organisms, oxIAA was nearly
omnipresent. Among green algae, ABA was only found in the culture
medium of charophytes that also contained it intracellularly. By con-
trast, land plants contained endogenous ABA but did not release it into
the culture media. The only cases of jasmonate content in the culture
media were represented by JA in the charophytes Interfilum para-
doxum and Coleochaete scutata, which also contained it intracellularly.
Further context to the measurements performed in biomass and
culture media is gained from the same analysis in blank media, which
contained no biological material. We revealed that IAA, ABA, jasmo-
nates, and most CK were absent in the blank media (black circles in
Figs. 1and 2b). Hence, their presence in either biomass or medium
resulted from the activity of living organisms. Ethylene release was
likewise detected only in the setups containing living material except
for one positive case in the blank medium of Chara braunii (Supple-
mentary Data 4), which might be explained as an artifact of the med-
ium composition. Certain other analytes were frequently detected in
the blank media. This includes oxIAA and iP-type CK, which were
present at lower levels in blanks compared to biomass (Fig. 1)andthe
culture media (Fig. 2b), suggesting that the living organisms addi-
tionally produced and released these compounds on their own. Phe-
nolics were likewise present in blanks, and the ratiotoward the content
in biomass and culture media varied by analyte and species. For
instance, Mougeotia scalaris and Mesotaenium endlicherianum pro-
duced PAA and released it into the environment. Klebsormidiophyceae
enriched the culture medium with PAA and BA but did not accumulate
these intracellularly. Selaginella uncinata degraded SA and BA present
in the medium upon inoculation, possibly by uptake and internal
metabolization. Summarily, we found that the detection of ABA, aux-
ins, CK, jasmonates and ethylene in either biomass or culture medium
could be entirely or predominantly attributed to the action of living
organisms. Phenolics were not only produced by the living biomass
but frequently also had been a part of the culture media prior to
inoculation with living cultures, and individual species differed in their
production, uptake, and release.
The content of certain phytohormones in blank media motivated
us to further address the possible sources of these contaminants by
analyzing different purity grades of water and three brands of agar
(Fig. 2c). Amongall analyzed compounds, the watersamples contained
only the phenolic BA. However, BA was the only analyte in our analysis
which lacked the qualifier ion, so the signal might not be entirely
specific to BA. The agar samples (incorporated in MilliQ grade water)
contained oxIAA, iP-type CK, and considerable amounts of the phe-
nolics BA, SA, and PAA (Fig. 2c, Supplementary Data 3). Hence, the low-
level detection of additional compounds in blank media compared to
these agar and water controls is likely to be attributed to different,
unknown sources of contamination.
Phytohormones and biological contamination
The issue of contamination of algal material by eukaryotic or prokar-
yotic microorganisms in phytohormone measurements and treat-
ments haslong been debated35,44. In our effort to obtain axenic cultures
whenever possible (see Supplementary Fig. 5 for the culture con-
tamination test), we managed to obtain three sets of charophyte
strains, corresponding to three charophyte lineages. Each set con-
tained two strains from the same lineage, one axenic and the other
contaminated. These sets constituted two strains of Mesostigma viride
(Mesostigmatophyceae), Chlorokybus cerffiiand C. melkonianii (both
Chlorokybophyceae), and two strains of Coleochaete scutata (both
Coleochaetophyceae). Remarkably, we found that phytohormone
profiles in the biomass were qualitatively similar within lineages,
regardless of strain contamination (Supplementary Fig. 6). Some
qualitative differences could be observed in the culture media, but
these were lineage-specific (Supplementary Fig. 6). For instance, the
contaminated Chlorokybus cerffiiculture medium contained IAA, in
contrast to the axenic strain Chlorokybus melkonianii.However,the
opposite pattern could be observed between the axenic and con-
taminated strains of Mesostigma viride and Coleochaete scutata.This
could be attributed to a lineage-specific reaction to contamination
among charophytes, or the activity of different (though unidentified)
microorganisms present in these cultures. In summary, we did not
detect any shared patterns in phytohormone biosynthesis and
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Nature Communications | (2024) 15:3875 4
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excretion in response to microbial contamination. This suggests that
lineage-specific patterns are likelymore profound than the influence of
microbial contamination.
Discussion
Comparing our results to the available literature on phytohormone
profiling, we see that certain compounds are being detected more
consistently than others. Our findings align with the ubiquitous
occurrence of IAA and oxIAA throughout Viridiplantae (Supplemen-
tary Data 5), much like the phenolic phytohormone SA5.IAA-amides
were absent in green algae in our screen and certain other studies25,26,
though isolated occurrences at low amounts had been reported
elsewhere18. The detection of the jasmonate JA had previously varied in
charophytes and bryophytes, while OPDA and dnOPDA were found to
ABA
IAA
IAM
oxIAA
tZ types
cZ types
iP types
JA
SA
PA A
BA
ABA AUXINS CYTOKININS JAs PHENOLICS
ABA
IAA
IAM
oxIAA
tZ types
cZ types
iP types
JA
SA
PA A
BA
MilliQ + agar #3
MilliQ + agar #2
MilliQ + agar #1
MilliQ (in flask)
MilliQ
dH2O
tap water
−2 02
concentration
[pmol/g or ml]
<1 1 10 100
ABA AUXINS CYTOKININS JAs PHENOLICS
n.d.
b
c
absent in
blank
−2 0 2
P. patens
(protonema)
P. patens
(gametophores)
M. polymorpha
Spirogyra sp.
M. scalaris
Closterium p-s-l
complex
M. endlicherianum
C. scutata
†Nitella sp.
C. braunii
K. nitens
I. paradoxum
C. melkonianii
M. viride
D. communis
C. sorokininana
T. annulata
P. salinarum
S. uncinata
(shoots)
S. uncinata
(rhizophores)
aMedium-Blank RelationMedium-Biomass Relation absent in
biomass
†) wild-collected (no blank)
detected only at stationary phase
MeS types
log(medium/biomass) log(medium/blank)
MeS types
Fig. 2 | Detection of phytohormone compounds in culture media and control
samples of water and agar. a Ratio between concentrations detected in culture
medium and the corresponding biomass, in logarithmic scale. Color code: blue
shading, compound prevalent in biomass; red shading, compound prevalent in
medium; black, compound absent in biomass. bRatio between concentrations
detectedin culture medium andblank medium (containing no biological material),
in logarithmic scale. Color code: blue shading, compound prevalent in blank; red
shading, compound prevalent in medium; black, compound absent in blank. The
circle sizein a,bdenotes the meanconcentration in culture media (pmolper gram
or ml). No circle, compound not detected (n.d.). Minimum n= 3 for culture media
(independent cultures; exact sample size for each strain is listed in Supplementary
Data 3), n= 2 for blank media.cCompounds detected in different purity grades of
water and three brands of agar (1.5% w/v, incorporated in MilliQ grade water). The
circle size denotes mean concentration (pmol per gram or ml); n=3samples.No
circle, compound not detected (n.d.). Compound abbreviations are listed in the
legend in Fig. 1. Data variation is shown in Supplementary Fig. 3 and listed in
Supplementary Data 8.
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be more consistent (Supplementary Data 5). We detected endogenous
jasmonates in multiple charophytes, often at considerable con-
centrations, but found land plants to be nearly devoid of jasmonates
under similar culture conditions. As suggested by the role of ODPA and
dnOPDA in a thermotolerance response conserved inStreptophyta15,45,
the jasmonate-rich charophyte strains in our analysis may have been
growing outside their temperature optimum. Additional stimuli such
as wounding regulate jasmonate production in land plants15, plausibly
accounting for their near-absence in land plant material under our
conditions. The published CK profiles in non-seed Viridiplantae have
been inconsistent at the lineage level. Despite numerous qualitative
disagreements (see Supplementary Data 5), our results at least support
two general trends in green algae and bryophytes: CK profiles are
dominated by the tRNA CK types, and CK-N-glucosides are absent18,19,46.
We detected ABA in all the tested land plants but only a few green
algae, whereas ABA presence in green algae and non-Viridiplantae
lineages has been well-established in literature17,26,27,42,47–49. Since ABA
was only found in stationary-phase chyrophyte samples, we speculate
that ABA production at detectable levels in green algae may result
from certain stress conditions that were rarely achieved among our
cultured strains. In summary, we confirm that free auxins, tRNA CK,
and SA are produced universally in green algae, bryophytes, and
lycophytes. By contrast, the biosynthesis of ABA in charophytes and
jasmonates in Streptophyta at detectable levels appears to depend on
certain stimuli, in our case manifested from culture conditions. We
found tZ and IAA- and CK-conjugates to be restricted to land plants,
but conflicting reports had been published. Regarding ethylene
release, to our knowledge, current literature lacks data comparable to
our dataset.
Our LC/MS profile in green algal and land plant biomass was
complemented with the same analysis performed in the correspond-
ing culture media. CK profiles in biomass were generally mirrored at
lower concentrations in culture media, supporting earlier isolated
reports of extracellular CK release in Viridiplantae50.Ithasbeen
reported that chlorophytes excrete ABA and IAA49,51–54, which is con-
trary to our findings. All chlorophytes in this study also lacked intra-
cellular ABA. On the other hand, charophytes that contained
intracellular ABA were also excreting it into the media. The excretion
of IAA in charophytes was previously proposed to reflect an ancestral
function in inter-organismal communication43,55.Wemeasured
increased IAA concentrations in the culture medium of Klebsormidium
nitens compared to biomass, coinciding with the conserved auxin
efflux function of the PIN-FORMED (PIN) proteins in the genus56.
However, we also found that cases of auxin excretion among char-
ophytes did not universally correlate with PIN conservation43.Inthe
control measurements of agar samples, the content of auxins, CK, and
SA may reflect the original biological source in agarophyte red
algae57–59. Although culturing plants on agar is equivalent to treatment
with ca. 10–20 nM SA, the effective doses used in treatments of plants
are much higher60. In all, we revealed that phytohormones were
released into the culture media in streptophytes. This mainly included
auxin, CK, and phenolics, with lineage-specific differences. Addition-
ally, we show that certain phytohormone compounds are present at
basal levels in culture media as contaminants from the media
components.
Our comparison of certain axenic vs. xenic (contaminated) char-
ophyte cultures revealed little influence of contaminants on the overall
phytohormone profiles. No conserved reaction to contamination was
revealed among three investigated charophyte lineages, in
either intracellular or extracellular phytohormone content. As for the
contaminating microorganisms, little interpretation could be made;
previous studies on phytohormone biosynthesis and responses in
bacteria and fungi have been focused dominantly on the strains
associated with vascular plants61–63. Although our results do not
suggest that the recognized phytohormones are relevant in
charophyte-microorganism interactions, a dedicated future investi-
gation should include the isolation and identification of microbial
strains living in natural association with the algae. Natural multi-algal
consortia could be studied in a similar fashion.
Plant evolution is researched in large part by tracking the con-
servation of genesknown from angiosperms.However, phytohormone
profiles in Viridiplantae regularly do not match genomic inferences.
IAA and oxIAA are omnipresent in Viridiplantae, whereas we only
detected glucosylated IAA in bryophytes and a lycophyte. Yet, the
enzymes recognized for auxin biosynthesis, non-decarboxylative IAA
oxidation, and IAA glucosylation, respectively, are only conserved in
land plants, angiosperms, and seed plants13. Inversely, the charophyte
Klebsormidium spp. lacks endogenous IAA amides (this study and
ref. 18) despite uniquely containing gene homologs of IAA-amide
synthases, which are otherwise restricted to land plants13. CK are well
known to regulate development in bryophytes via signaling conserved
with angiosperms64, but no such knowledge exists in charophytes.
Correspondingly,wefoundonlylandplantsamongthesampled
organisms to contain tZ-type CKs, which are known to be highly
bioactive65. However, these investigated land plants lack the adenylate-
type isopentenyltransferases recognized for tZ biosynthesis, i.e., do
not differ from green algae in the genomic evidence for CK
biosynthesis12. To make matters more complicated, green algae can
process applied IAA into IAA-amides and break down applied tZinto
adenine and adenosine, despite lacking IAA-amidases and CK
dehydrogenases18,23. We found all but one of the investigated Vir-
idiplantae to release ethylene into the environment. Yet, the enzyme
directly responsiblefor ethylene biosynthesis is only conserved in seed
plants14. Two SA biosynthesis pathways are deeply conserved in land
plants. Among green algae, only certain charophyte lineages contain
all the known components of at least one (the β-oxidation-based) of
these pathways5. Regardless, SA is produced ubiquitously in Vir-
idiplantae (our study and5). Interestingly, the chlorophyte Chlamydo-
monas reinhardtii lacks some genes of the β-oxidation-based SA
biosynthesis pathway, but a mutation in one enzyme conserved with
the angiosperm Oryza sativa results in decreased endogenous SA
levels in both species5. These examples remind us there is much left
unknown in the lineages outside of seed plants, including secondary
metabolism6. More research into these organisms will help us over-
come the limits of viewing plant evolution through the prism of
knowledge gained mainly in angiosperms.
The question of the origins of phytohormone function has been
approached by analytical profiling of native compounds, treatments
by externally applied phytohormones, and comparative genomics.
Biotechnology-motivated studies have revealed that unicellular
chlorophytes natively produce the most known phytohormones and
show growth responses to their external application. Similar results
were obtained in “microalgae”from non-Viridiplantae lineages,
including Eustigmatophyceae, Euglenozoa, or Cyanobacteria
(reviewed in refs.19,66,67). As phytohormones are defined as “natural
compounds affecting physiological processes at low concentrations“3,
these observations would suggest that phytohormone identity of
multiple organic compounds is widespread in the tree of life. However,
no underlying mechanism, genetic or otherwise, conserved between
land plants and any of these lineages has yet been shown. Research
through comparative genomics provides a different perspective; the
molecular machineries underlying the responses to most phyto-
hormones only became operational in the last common ancestor of
land plants or later in this lineage4–9. Our results showed that land
plants but not green algae (chlorophytes or charophytes) consistently
contained ABA and produced auxinand CK metabolites. Otherstudies
focusing on these hormones have likewise found differences between
land plants and green algae. In treatments, lower micromolar doses of
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 6
Content courtesy of Springer Nature, terms of use apply. Rights reserved

ABA, auxin, and CK elicit growth phenotypes in land plants such as
bryophytes68–72, but comparably higher doses of auxin and ABA (and
much higher than the levels detected as endogenous in this study) are
necessary to achieve comparably milder effects in charophytes73–76.No
data are available regarding CK treatments in charophytes. Compara-
tive genomics revealed that the genes coding for auxin and ABA
receptors are functionally conserved across land plants but are absent
in charophytes71,72,77,78, whereas the CK receptor homologs present in
charophytes arerelated to a cladethat does not act in CK perception in
land plants12. Additionally, gene homologs of CK signaling in the
charophyte Spirogyra pratensis are not transcriptionally responsive to
applied CK79. Collectively, our results together with the above-listed
observations imply that ABA, auxin, and CK might only have assumed
phytohormoneidentity in an ancestralland plant. Itmust be noted that
much morework lies ahead of us to properly understand the evolution
of phytohormone responses. Recent research has revealed that both
charophytes and land plants utilize OPDA and dnOPDA in a thermo-
tolerance response, which is distinct from the role of these compounds
in jasmonate signaling that only evolved in land plants15,45.Canonical
auxin signaling is likewise a land plant invention77,yetappliedIAA
elicits rapid changes in plasma membrane potential and protein
phosphorylation in both land plants and charophytes; while the native
function in charophytes remains unknown, it may involve a reaction to
light or osmotic stress80. As charophytes become better explored and
established as model organisms, we are likely to be updating our views
regarding the evolution and origins of phytohormone responses.
How can we interpret the presence of phytohormones in organ-
isms that lack a bona fide phytohormone response? The similar pro-
files of IAA and tRNA CK in many green, red, and brown algae, fungi,
animals, and other organisms17,18,48,57,81–83 might simply reflect the
metabolism of indoles and purines/tRNA84,85. Similarly, the presence of
ABA in algae such as Dunaliella spp. may be a side-effect of carotenoid
metabolism22,86,87. Increased ABA production from carotenoid oxida-
tion under stress conditions has been proposedas an evolutionary pre-
requisite to ABA functioning as a stress hormone6.Indeed,extant
green algae can accumulate ABA under stress without signs of ABA
itself affecting stress responses88. On a related note, it may be worth
considering that the compounds recognized as phytohormones, such
as IAA or CK, might only reflect phytohormone action when natively
present in amounts that exceed specific thresholds. After all, phyto-
hormone function in angiosperms follows a strict dose-response
relationship and is often coupled with their considerable accumulation
in specific tissues and developmental stages89–92. Therefore, if com-
pounds known as phytohormones have physiological functions in
extant charophytes, conserved with land plants or not, these may be
spatiotemporally or developmentally restricted (e.g., to developing
zygospores). Uncovering such possible functions would yet require a
considerable amount of research.
In the past two decades, bryophytes have been the torch illu-
minating the dark corners of plant evolutionary research. A new
frontier is currently opening in charophyte algae, perhaps even
more challenging due to longer divergence times coupled with
considerable diversity in morphology and physiology. We revealed
that charophytes are capable of producing most organic com-
pounds known as active phytohormones in land plants. However,
certain patterns in biosynthesis and metabolism coupled with
genomic and empirical evidence indicate that major, perhaps fun-
damental innovations underlying phytohormone action have only
evolved after the transition to land. Hence, this study represents
one piece in the puzzle of the origins and evolution of phyto-
hormone responses.
Methods
Chemicals
Supplementary data 6 lists the chemicals used in this study.
Algal and plant strains and cultivation
Algal strains were obtained from the following institutions: Microbial
culture collection, National Institute for Environmental Studies, Tsu-
kuba, Japan (NIES); Culture collection of algae, Goettingen University,
Germany (SAG); Central collection of algal cultures, Duisburg Uni-
versity, Essen, Germany (CCAC); Culture collection of algae, University
of Texas at Austin, USA (UTEX); Culture collection of algae, Depart-
ment of Botany, Charles University, Prague, Czechia (CAUP). Selagi-
nella uncinata (“Comenius”, original strain) was obtained from the
Botanical garden at the Comenius University, Bratislava, Slovakia, and
established asa sterile culture. The wild-collected samples of Nitella sp.
and the surrounding water (flash-frozen in liquid N
2
in situ) were
obtained from a freshwater spring pond under sandstone rocks (GPS:
50.6402767 N, 14.5147078E) in April 2021; the surrounding water was
notably pure and the algae not obviously covered with epiphytic
microflora.
See Supplementary Data 1 for a complete list of strains with source
identifiers and the culture media used. Murashige–Skoog medium
(Duchefa M0221), Gamborg medium (Duchefa G0210), and Bold’s
Basal Medium (BBM; Merck B5285) were purchased commercially.
BCD medium supplemented with diammonium tartrate (BCDAT) was
prepared according to93. C medium, Pro medium, and a modified
version of the SWCN-4 medium for Charophyceae were prepared per
instructions from the NIES collection (https://mcc.nies.go.jp/
02medium-e.html). Modified SWCN-4 as follows: garden soil was
mixed with river sand (1:4), dampened with dH
2
O, autoclaved 3×, laid
into sterile test tubes (3 cm height), and supplemented with 40 ml
sterile dH
2
O. BBM, C, and Pro media were supplemented with custo-
mized vitamin doses: B
12
10 mg/l medium, B
7
2mg/l, B
1
10 mg/l. Indi-
vidual vitamin stocks (1000×) were dissolved in dH
2
O, sterilized by
filtration (Millex SLGS033SS), stored at −20 °C, and added into cooled-
down but not yet solid agar-supplemented medium, or upon algal
inoculation (liquid medium). All media were sugar-free.
Land plants were cultured on solid media in plates sealed with
surgical tape (Micropore 1530‑0). Marchantia polymorpha were
inoculated from individual gemmae. Physcomitrium patens protonema
were homogenized weekly using Ika T25 Digital Ultra Turrax with 8G
disperser tool (IKA-Werke, Germany) and spread on plates containing
medium overlaid with cellophane foil. When necessary, cultures were
left to grow without homogenization to allow gametophore emer-
gence. Selaginella uncincata (chosen for itssuperb growthin vitro) was
inoculated from apical branch cuttings. 2–3 apical explants of Chara
braunii (each with at least two nodes) were inoculated per one test
tube containing soil and culture medium. Other algae, if not grown for
analysis, were inoculated into the fresh medium from a fraction of the
original culture (2–10% biomass) in 6–8 week intervals or longer,
depending on the growth character of each strain.
Although we have generally avoided this measure in cultures
intended for phytohormone analysis, certain algal strains (Supple-
mentary Data 1) require supplementation of the c ulture media with soil
extract for continuous proper growth. Soil extract was prepared by
mixing soil with dH
2
O (1:3 volume), autoclaving, passing through filter
paper overnight, and autoclaving again. The soil (without obvious leaf
litter) was collected in a submontane forest of European beech (Fagus
sylvatica; GPS: 50.8653578 N, 15.1046144E) at the end of winter.
All living material was cultured at 23 °C, 16:8 hours light:dark
regime. The cultures were illuminated by mixed-spectrum fluorescent
light tubes using Osram Fluora TLD 36 W (Osram Licht AG, Germany)
and Philips Master TLD Super 36 W (Koninklijke Philips N.V., The
Netherlands); see Supplementary Data 7 for details on illumination
spectra and intensity.
Culture contamination tests
The axenic status of living cultures was examined by two approaches.
First, the algal biomass used for analysis was observed under a
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 7
Content courtesy of Springer Nature, terms of use apply. Rights reserved

microscope (Supplementary Fig. 5). Second, the algal and land plant
biomass was inoculated on four types of media designed to support
the growth of bacteria and fungi. These media were labeled as LB,
Euglena, Trebouxia, and Sabouraud (all supplemented with 1.5% w/v
agar, Duchefa P1001) and were prepared per instructions of the Culture
Collection of Algae at the University of Texas, Austin (UTEX) (https://
utex.freshdesk.com/support/solutions/articles/19000006667-how-do-
you-test-for-contamination-). After inoculation, the plates were sealed
with parafilm and cultured in the dark at 37 °C for 7 days. All plates
were photographed and examined microscopically (Supplemen-
tary Fig. 5).
Sample preparation
Algal biomass was sampled during the proliferative phase of culture
growth (1.5–3 weeks after inoculation, depending on individual
strains). Streptophyte algae were additionally sampled during the
stationary phase of growth (4–6weeks).Marchantia polymorpha thalli
were sampled on day 19. Physcomitrium patens protonema and
gametophores were sampled after 1 and 5 weeks of growth, respec-
tively. Selaginella uncinata was sampled after 5 weeks of growth.
Samples for LC/MS were harvested as follows (see Supplemen-
trary Data 1): liquid-cultured algae were collected by pipetting (cutoff
tips) and filtration through a nylon mesh filter (20 μm, Merck Millipore
NY2004700) using underpressure. Strains with smaller cells, i.e.,
Picocystis salinarum,Desmodesmus communis,Mesostigma viride,and
Closterium p-s-l complex were collected by centrifugation (1000 × g)
and supernatant decantation. Algae and moss protonema growing on
solid media were scraped with a spatula and filtered to remove
excessive moisture. Chara braunii and Nitella sp. thalli (without rhi-
zoids) were washed with dH
2
O, and excessive moisture was removed
with a cotton pad. Bryophyte thalli and Selaginella uncinata apical
cuttings were directly transferred into collection tubes. Biomass
samples (10-50 mg fresh weight each) were transferred into 2-ml thick-
walled microcentrifuge tubes (SSIbio 2340-00, Scientific Specialities,
USA) with screwable lids (SSIbio 2002-00), flash-frozen in liquid
nitrogen and stored at −80 °C. Liquid culture media were sampled by
pipetting, centrifugation (1000 × g), and decanting as 100 µl samples.
Samples of solid culture media were scraped from inside the solidagar
(avoiding any residual biomass) with a cut pipette tip and sampled as
ca. 50 mg each. Blank media were sampled similarly. The media sam-
ples were likewise flash-frozen in liquid nitrogen and stored at −80 °C.
Phytohormone analysis
Samples were extracted with 100 µl 1 M formic acid solution. The fol-
lowing isotope-labelled standards were added at 1 pmol per sample:
13C
6
-IAA (Cambridge Isotope Laboratories, Tewksbury, MA, USA); 2H
4
-
SA (Sigma-Aldrich, St. Louis, MO, USA); 2H
3
-PA, 2H
3
-DPA (NRC-PBI);
2H
6
-ABA, 2H
5
-JA, 2H
5
-tZ, 2H
5
-tZR, 2H
5
-tZRMP, 2H
5
-tZ7G, 2H
5
-tZ9G, 2H
5
-
tZOG, 2H
5
-tZROG, 15N
4
-cZ, 2H
3
-DZ, 2H
3
-DZR, 2H
3
-DZ9G, 2H
3
-DZRMP,
2H
7
-DZOG, 2H
6
-iP, 2H
6
-iPR, 2H
6
-iP7G, 2H
6
-iP9G, 2H
6
-iPRMP 2H
2
-GA19,
(2H
5
)(15N
1
)-IAA-Asp and (2H
5
)(15N
1
)-IAA-Glu (Olchemim, Olomouc,
Czech Republic). The extracts were centrifuged at 30,000 × gat 4 °C.
The supernatants were applied to SPE Oasis HLB 96-well column plates
(10 mg/well; Waters, Milford, MA, USA) conditioned with 100 µLacet-
onitrile and 100 µl 1 M formic acid using Pressure+ 96 manifold (Bio-
tage, Uppsala, Sweden). After washing the wells three times with 100 µl
water, the samples were eluted with 100 µl 50% acetonitrile in water.
The eluates were separated on Kinetex EVO C18 HPLC column (2.6 µm,
150 × 2.1 mm, Phenomenex, Torrance, CA, USA). Mobile phases were as
follows: A, 5 mM ammonium acetate, and 2 µM medronic acid in water;
B, 95:5 acetonitrile:water (v/v). The following gradient was applied: 5%
B in 0 min, 5-7% B (0.1–5 min), 10-35% B (5.1–12 min), and 35-100% B (12-
13 min), followed by a 1 min hold at100% B (13-14 min) and return to 5%
B. Hormone analysis was performed with an LC/MS system consisting
of UHPLC 1290 Infinity II (Agilent, Santa Clara, CA, USA) coupled to
6495 Triple Quadrupole Mass Spectrometer (Agilent, Santa Clara, CA,
USA). The Jet Stream (AJS) ion source parameters included: gas tem-
perature 180 °C, gas flow 19 l/min, sheath gas temperature 400 °C,
sheath gas flow 12 l/min, nebulizer pressure 25 psi, capillary voltage:
positive/negative −3000 V/2500 V, nozzle voltage: positive/negative
−0 V/1000 V. The compounds were analysed in multiple reaction
monitoring modes (transitions are listed in Supplementary Data 2),
with quantification by the isotope dilution method. Data acquisition
and processing were performed with Mass Hunter software B.08
(Agilent, Santa Clara, CA, USA). Data processing was performed using
Agile2 integration algorithm followed by manual verification. For cal-
culation of concentrations reflecting the amount of sample and
internal standards, see Supplementary Data 3.
For ethylene measurements, we adapted an established
protocol94.Blankmediawereprocessedfirst to determine any possible
background levels or noise and enable subsequent correction of the
biomass samples. A set volume (Supplementary Data 4) of liquid or
solid medium in clear-glass 10-ml chromatography vials (Thermo
Fisher 10623633) was left standing (lid open) for 1 hour. The vials were
then sealed air-tight with a rubber septum (Thermo Fisher 11845060)
and a snap cap (Thermo Fisher 11805020) using a manual crimping
tool (Thermo Fisher 10372525), then kept at 21 °C in low light condi-
tions (35 µmol m−2s−1)under4fluorescent tubes (Philips TLD 36 W/33-
640). After 6 hours, ethylene levels within the headspace were mea-
sured using laser-based photoacoustic spectroscopy (ETD-300, Sensor
Sense) in stop-and-flow mode. Algal biomass samples were suspended
in fresh medium (same volume as used in blank measurements, see
Supplementary Data 4) and transferred into the chromatography vials.
Land plant samples were transferred into chromatography vials con-
taining solid culture medium and a minimum volume of pure water
(MilliQ grade) (Supplementary Data 4); no pure water was added to
Physcomitrium patens gametophores, which were previously cultured
in the vials. The biomass samples were then processed as the blank
media, i.e., left open to acclimatize for 1 h, sealed for 6 h, and measured
for total ethylene accumulated. Samples were measured in 5 biological
and 5 blank medium replicates. The system was calibrated using
standard ethylene mixtures (Air Liquide 23209012).
Presentation of results and statistical evaluation
Phytohormone profiles are presented as means and related to other
sample categories by logarithmic ratio to visualize possible back-
ground of compounds in media (relation to corresponding blank
media), or possible excretion of compounds to media (relation to
culture media). Relative errors of biological and technical replications
of LC/MS analysis are provided in Supplementary Fig. 3 and Supple-
mentary Data 8. The significance of cultivation effects (e.g., culture
stage and strainaxenicity; Supplementary Figs. 2 and 6)were evaluated
by linear mixed effects model after logarithmic transformation (lme4
package95) and tested by Likelihood-ratio test (‘lmerTest’package96.
Differences between groups were determined by post-hoc comparison
(Tukey method, ‘emmeans’package97. The same statistical evaluation
was used to determine the significance of ethylene emanation by the
biomass, due to the presence of mediaduring ethylene measurements
on living material (Supplementary Fig. 4). Statistical comparisons of
LC/MS data between biomass/culture media and blank media were not
performed, as these were analyzed as separate samples. Statistical
analyses and data plotting were performed using R software package
4.2.3 (R Core Team) with ‘tidyverse’package environment (v2.0.0).
Figures were finally composed using Adobe Illustrator 2020.
Reporting summary
Further information on research design is available in the Nature
Portfolio Reporting Summary linked to this article.
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 8
Content courtesy of Springer Nature, terms of use apply. Rights reserved

Data availability
All raw LC/MS data generated in this study have been deposited in the
Zenodo repository: https://doi.org/10.5281/zenodo.10411071.Source
data are provided with this paper.
Code availability
Code for data plotting and statistical analyses is deposited at the
following links: https://github.com/VoSchmidt/BubblePlots-for-
phytohormones https://github.com/vosolsob/phytohormone.
References
1. Bowles,A.M.C.,Williamson,C.J.,Williams,T.A.,Lenton,T.M.&
Donoghue, P. C. J. The origin and early evolution of plants. Trends
Plant Sci. 28,312–329 (2023).
2. Delwiche,C.F.Thegenomesof charophyte green algae. Adv. Bot.
Res.78,255–270 (2016).
3. Davies, P. J. The plant hormones: their nature, occurrence, and
functions. In: Plant hormones 1–15 (Springer Netherlands, 2010).
https://doi.org/10.1007/978-1-4020-2686-7_1.
4. Bowman,J.L.,Briginshaw,L.N.,Fisher,T.J.&Flores-Sandoval,E.
Something ancient and something neofunctionalized—evolution of
land plant hormone signaling pathways. Curr. Opin. Plant Biol. 47,
64–72 (2019).
5. Jia, X. et al. The origin and evolution of salicylic acid signaling and
biosynthesis in plants. Mol. Plant 16,245–259 (2023).
6. Rieseberg, T. P. et al. Crossroads in the evolution of plant specia-
lized metabolism. Semin. Cell Dev. Biol. 134,37–58 (2023).
7. Hernández-García, J., Briones-Moreno, A. & Blázquez, M. A. Origin
and evolution of gibberellin signaling and metabolism in plants.
Semin. Cell Dev. Biol. 109,46–54 (2021).
8. Kyozuka, J., Nomura, T. & Shimamura, M. Origins and evolution of
the dual functions of strigolactones as rhizosphere signaling
molecules and plant hormones. Curr. Opin. Plant Biol. 65,
102154 (2022).
9. Zheng, B. et al. Evolutionary analysis and functional identification of
ancient brassinosteroid receptors in Ceratopteris richardii. Int. J.
Mol. Sci. 23, 6795 (2022).
10. Dabravolski, S. A. & Isayenkov, S. V. Evolution of the cytokinin
dehydrogenase (CKX) domain. J. Mol. Evol. 89,665–677 (2021).
11. Chen, L., Jameson, G. B., Guo, Y., Song, J. & Jameson, P. E. The
LONELY GUY gene family: from mosses to wheat, the key to the
formation of active cytokinins in plants. Plant Biotechnol. J. 20,
625–645 (2022).
12. Powell, A. E. & Heyl, A. The origin and early evolution of cytokinin
signaling. Front. Plant Sci. 14, 1142748 (2023).
13. Bowman, J. L., Flores Sandoval, E. & Kato, H. On the evolutionary
origins of land plant auxin biology. Cold Spring Harb. Perspect. Biol.
13, a040048 (2021).
14. Li, D., Mou, W., Van de Poel, B. & Chang, C. Something old, some-
thing new: conservation of the ethylene precursor 1-amino-cyclo-
propane-1-carboxylic acid as a signaling molecule. Curr. Opin. Plant
Biol. 65, 102116 (2022).
15. Chini, A., Monte, I., Zamarreño, A. M., García‐Mina,J.M.&Solano,R.
Evolution of the jasmonate ligands and their biosynthetic pathways.
N. Phytol. 238, 2236–2246 (2023).
16. Tsa vkelova, E. A., Klimova, S. Y., Cherdyntseva, T. A. & Netrusov, A. I.
Hormones and hormone-like substances of microorganisms: a
review. Appl. Biochem. Microbiol. 42, 229–235 (2006).
17. Morrison, E. N. et al. Detection of phytohormones in temperate
forest fungi predicts consistent abscisic acid production and a
common pathway for cytokinin biosynthesis. Mycologia 107,
245–257 (2015).
18. Žižková, E. et al. Control of cytokinin and auxin homeostasis in
cyanobacteria and algae. Ann. Bot. 119,151–166 (2017).
19. Stirk, W. A. & van Staden, J. Potential of phytohormones as a strat-
egy to improve microalgae productivity for biotechnological
applications. Biotechnol. Adv. 44,107612(2020).
20. Koeduka, T. et al. Biochemical characterization of allene oxide
synthases from the liverwort Marchantia polymorpha and green
microalgae Klebsormidium flaccidum provides insight into the
evolutionary divergence of the plant CYP74 family. Planta 242,
1175–1186 (2015).
21. Gachet, M. S., Schubert, A., Calarco, S., Boccard, J. & Gertsch, J.
Targeted metabolomics shows plasticity in the evolution of sig-
naling lipids and uncovers old and new endocannabinoids in the
plant kingdom. Sci. Rep. 7, 41177 (2017).
22. Tietz, A., Ruttkowski, U., Kohler, R. & Kasprik, W. Further investiga-
tions on the occurrence and the effects of abscisic acid in algae.
Biochem. Physiol. Pflanz. 184,259–266 (1989).
23. Sztein, A. E., Cohen, J. D. & Cooke, T. J. Evolutionary patterns in the
auxinmetabolismofgreenplants.Int. J. Plant Sci. 161,849–859
(2000).
24. Delaux, P. M. et al. Origin of strigolactones in the green lineage. N.
Phytol. 195,857–871 (2012).
25. Stirk, W. A. et al. Auxin and cytokinin relationships in 24 microalgal
strains. J. Phycol. 49,459–467 (2013).
26. Hackenberg, D. & Pandey, S. Heterotrimeric G-proteins in green
algae. Plant Signal. Behav. 9,e28457(2014).
27. Hori, K. et al. Klebsormidium flaccidum genome reveals primary
factors for plant terrestrial adaptation. Nat. Commun. 5,
3978 (2014).
28. Beilby,M.J.,Turi,C.E.,Baker,T.C.,Tymm,F.J.M.&Murch,S.J.
Circadian changes in endogenous concentrations of indole-3-
acetic acid, melatonin, serotonin, abscisic acid and jasmonic acid in
Characeae (Chara australis Brown). Plant Signal. Behav. 10,
e1082697 (2015).
29. Ju, C. et al. Conservation of ethylene as a plant hormone over 450
million years of evolution. Nat. Plants 1,1–24 (2015).
30. Tietz,A.&Kasprik,W.Identification of abscisic acid in a green alga.
Biochem. Physiol. Pflanz. 181,269–274 (1986).
31. Yamamoto, Y. et al. Functional analysis of allene oxide cyclase,
MpAOC, in the liverwort Marchantia polymorpha. Phytochemistry
116,48–56 (2015).
32. Monte, I. et al. Ligand-receptor co-evolution shaped the jasmonate
pathway in land plants. Nat. Chem. Biol. 14,480–488 (2018).
33. Wang, S. et al. G enomes of early-diverging streptophyte algae shed
light on plant terrestrialization. Nat. Plants 6,95–106 (2020).
34. Haig, D. Coleochaete and the origin of sporophytes. Am. J. Bot. 102,
417–422 (2015).
35. Cook, S. D. An historical review of phenylacetic acid. Plant Cell
Physiol. 60,243–254 (2019).
36. Widhalm,J.R.&Dudareva,N.Afamiliar ring to it: biosynthesis of
plant benzoic acids. Mol. Plant 8,83–97 (2015).
37. Kaźmierczak, A. & Stepiński, D. GA3 content in young and mature
antheridia of Chara tomentosa estimated by capillary electrophor-
esis. Folia Histochem. Cytobiol. 43,65–67 (2005).
38. Hirano, K. et al. The GID1-mediated gibberellin perception
mechanism is conserved in the lycophyte selaginella moellendorffii
but not in the bryophyte physcomitrella patens. Plant Cell 19,
3058–3079 (2007).
39. Stirk, W. A. et al. Hormone profiles in microalgae: Gibberellins and
brassinosteroids. Plant Physiol. Biochem. 70,348–353 (2013).
40. Bajguz, A. Isolation and characterization of brassinosteroids from
algal cultures of Chlorella vulgaris Beijerinck (Trebouxiophyceae). J.
Plant Physiol. 166, 1946–1949 (2009).
41. Proust, H. et al. Strigolactones regulate protonema branching and
act as a quorum sensing-like signal in the moss Physcomitrella
patens. Development 138,1531–1539 (2011).
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 9
Content courtesy of Springer Nature, terms of use apply. Rights reserved

42. Hartung, W. The evolution of abscisic acid (ABA) and ABA function
in lower plants, fungi and lichen. Funct. Plant Biol. 37, 806 (2010).
43. Vosolsobě,S.,Skokan,R.&Petrášek, J. The evolutionary origins of
auxin transport: what we know and what we need to know. J. Exp.
Bot. 71,3287–3295 (2020).
44. Evans, L. V. & Trewavas, A. J. Is algal development controlled by
plant growth substances? J. Phycol. 27,322–326 (1991).
45. Monte, I. et al. An ancient COI1-independent function for reactive
electrophilic oxylipins in thermotolerance. Curr. Biol. 30,
962–971.e3 (2020).
46. Záveská Drábková, L., Dobrev, P. I. & Motyka, V. Phytohormone
profiling across the bryophytes. PLoS One 10, e0125411 (2015).
47. Lu, Y. et al. Antagonistic roles of abscisic acid and cytokinin during
response to nitrogen depletion in oleaginous microalga N anno-
chloropsis oceanica expand the evolutionary breadth of phyto-
hormone function. Plant J. 80,52–68 (2014).
48. Noble,A.,Kisiala,A.,Galer,A.,Clysdale,D.&Emery,R.J.N.Euglena
gracilis (Euglenophyceae) produces abscisic acid and cytokinins
and responds to their exogenous application singly and in combi-
nation with other growth regulators. Eur. J. Phycol. 49,
244–254 (2014).
49. Pichler, G. et al. Abundance and extracellular release of phyto-
hormones in aero‐terrestrial microalgae (Trebouxiophyceae,
Chlorophyta) as a potential chemical signaling source 1. J. Phycol.
56,1295–1307 (2020).
50. Stirk, W. A. & van Staden, J. Flow of cytokinins through the envir-
onment. Plant Growth Regul. 62,101–116 (2010).
51. Maršálek, B., Zahradníčková, H. & Hronková, M. Extracellular pro-
duction of abscisic acid by soil algae under salt, acid or drought
stress. Z.f.ür.Naturforsch.C.47,701–704 (1992).
52. Mazur, H., Konop, A. & Synak, R. Indole-3-acetic acid in the culture
medium of two axenic green microalgae. J. Appl. Phycol. 13,
35–42 (2001).
53. Prieto,C.R.E.,Cordoba,C.N.M.,Montenegro,J.A.M.&González-
Mariño, G. E. Production of indole-3-acetic acid in the culture
medium of microalga Scenedesmus obliquus (UTEX 393). J. Braz.
Chem. Soc. 22,2355–2361 (2011).
54. Khasin, M., Cahoon, R. R., Nickerson, K. W. & Riekhof, W. R. Mole-
cular machinery of auxin synthesis, secretion, and perception in the
unicellular chlorophyte alga Chlorella sorokiniana UTEX 1230. PLoS
One 13, e0205227 (2018).
55. Bennett, T. PIN proteins and the evolution of plant development.
Trends Plant Sci. 20,498–507 (2015).
56. Skokan, R. et al. PIN-driven auxin transport emerged early in
streptophyte evolution. Nat. Plants 5, 1114–1119 (2019).
57. Yokoya, N. S. et al. Endogenous cytokinins, auxins, and abscisic acid
in red algae from brazil 1. J. Phycol. 46, 1198–1205 (2010).
58. Sabeena Farvin, K. H. & Jacobsen, C. Phenolic compounds and
antioxidant activities of selected species of seaweeds from Danish
coast. Food Chem. 138,1670–1681 (2013).
59. Hou, S., Lin, L., Lv, Y., Xu, N. & Sun, X. Responses of lipoxygenase,
jasmonic acid, and salicylic acid to temperature and exogenous
phytohormone treatments in Gracilariopsis lemaneiformis (Rhodo-
phyta). J. Appl. Phycol. 30, 3387–3394 (2018).
60. Beyer, S. F. et al. Disclosure of salicylic acid and jasmonic acid-
responsive genes provides a molecular tool for deciphering stress
responses in soybean. Sci. Rep. 11, 20600 (2021).
61. Spaepen, S. & Vanderleyden, J. Auxin and plant-microbe interac-
tions. Cold Spring Harb. Perspect. Biol. 3,a001438–a001438 (2011).
62. Lievens, L., Pollier, J., Goossens, A., Beyaert, R. & Staal, J. Abscisic
acid as pathogen effector and immune regulator. Front. Plant Sci. 8,
587 (2017).
63. Frébortová, J. & Frébort, I. Biochemical and structural aspects of
cytokinin biosynthesis and degradation in bacteria. Microorganisms
9,1314(2021).
64. Rashotte, A. M. The evolution of cytokinin signaling and its role in
development before Angiosperms. Semin. Cell Dev. Biol. 109,
31–38 (2021).
65. Hluska, T., Hlusková, L. & Emery, R. J. N. The hulks and the dead-
pools of the cytokinin universe: a dual strategy for cytokinin pro-
duction, translocation, and signal transduction. Biomolecules 11,
209 (2021).
66. Lu, Y. & Xu, J. Phytohormones in microalgae: a new opportunity for
microalgal biotechnology? Trends Plant Sci. 20,273–282 (2015).
67. Han, X., Zeng, H., Bartocci, P., Fantozzi, F. & Yan, Y. Phytohormones
and effects on growth and metabolites of microalgae: a review.
Fermentation 4,25(2018).
68. Ashton, N. W., Grimsley, N. H. & Cove, D. J. Analysis of gameto-
phytic development in the moss, Physcomitrella patens, using
auxin and cytokinin resistant mutants. Planta 144,427–435 (1979).
69. Yasumura, Y., Crumpton-Taylor, M., Fuentes, S. & Harberd, N. P.
Step-by-step acquisition of the gibberellin-DELLA growth-reg-
ulatory mechanism during land-plant evolution. Curr. Biol. 17,
1225–1230 (2007).
70. Lavy, M. et al. Constitutive auxin response in Physcomitrella reveals
complex interactions between Aux/IAA and ARF proteins. Elife 5,
e13325 (2016).
71. Jahan, A. et al. Archetypal roles of an abscisic acid receptor in
drought and sugar responses in liverworts. Plant Physiol. 179,
317–328 (2019).
72. Suzuki, H., Kato, H., Iwano, M., Nishihama, R. & Kohchi, T. Auxin sig-
naling is essential for organogenesis but not for cell survival in the
liverwort Marchantia polymorpha. Plant Cell 35,1058–1075 (2023).
73. Klambt, D., Knauth, B. & Dittmann, I. Auxin dependent growth of
rhizoids of Chara globularis. Physiol. Plant. 85,537–540 (1992).
74. Sederias, J. & Colman, B. The interaction of light and low tem-
perature on breaking the dormancy of Chara vulgaris oospores.
Aquat. Bot. 87, 229–234 (2007).
75. Nagao,M.,Matsui,K.&Uemura,M.Klebsormidiumflaccidum, a
charophycean green alga, exhibits cold acclimation that is closely
associated with compatible solute accumulation and ultra-
structural changes. Plant. Cell Environ. 31,872–885 (2008).
76. Ohtaka,K.,Hori,K.,Kanno,Y.,Seo,M.&Ohta,H.Primitiveauxin
response without TIR1 and Aux/IAA in the charophyte alga Kleb-
sormidium nitens. Plant Physiol. 174,1621–1632 (2017).
77. Mutte, S. K. et al. Origin and evolution of the nuclear auxin response
system. Elife 7,25(2018).
78. Sun, Y. et al. A ligand-independent origin of abscisic acid percep-
tion. Proc.NatlAcad.Sci.116,24892–24899 (2019).
79. de Vr ies, J. et al. Heat stress response in the closest algal relatives of
land plants reveals conserved stress signaling circuits. Plant J. 103,
1025–1048 (2020).
80. Kuhn, A. et al. RAF-like protein kinases mediate a deeply conserved,
rapid auxin response. Cell 187,130–148.e17 (2024).
81. Le Bail, A. et al. Auxin metabolism and function in the multicellular
brown alga Ectocarpus siliculosus. Plant Physiol. 153,128–144 (2010).
82. Seegobin, M. et al. Canis familiaris tissues are characterized by
different profiles of cytokinins typical of the tRNA degradation
pathway. FASEB J. 32,6575–6581 (2018).
83. Aoki, M. M. et al. Phytohormone metabolism in human cells: cyto-
kinins are taken up and interconverted in HeLa cell culture. FASEB
BioAdv. 1,320–331 (2019).
84. Gibb,M.,Kisiala,A.B.,Morrison,E.N.&Emery,R.J.N.Theorigins
and roles of methylthiolated cytokinins: evidence from among life
kingdoms. Front. Cell Dev. Biol. 8,605672(2020).
85. Liu, H. et al. Non-target metabolomics reveals the changes of small
molecular substances in duck breast meat under different pre-
servation time. Food Res. Int. 161, 111859 (2022).
86. Hirsch, R., Hartung, W. & Gimmler, H. Abscisic acid content of algae
under stress. Bot. Acta 102,326–334 (1989).
Article https://doi.org/10.1038/s41467-024-47753-z
Nature Communications | (2024) 15:3875 10
Content courtesy of Springer Nature, terms of use apply. Rights reserved

87. Xu, Y. & Harvey, P. J. Carotenoid production by dunaliella salina
under red light. Antioxidants 8,123(2019).
88. Sun, Y., Pri-Tal, O., Michaeli, D. & Mosquna, A. Evolution of abscisic
acid signaling module and its perception. Front. Plant Sci. 11,
934 (2020).
89. Müller, A., Düchting, P. & Weiler, E. A multiplex GC-MS/MS tech-
nique for the sensitive and quantitative single-run analysis of acidic
phytohormones and related compounds, and its application to
Arabidopsis thaliana. Planta 216,44–56 (2002).
90. Dobrev, P. I., Havlíček, L., Vágner, M., Malbeck, J. & Kamínek, M.
Purification and determination of plant hormones auxin and absci-
sic acid using solid phase extraction and two-dimensional high
performance liquid chromatography. J. Chromatogr. A 1075,
159–166 (2005).
91. Skalák, J. et al. Multifaceted activity of cytokinin in leaf develop-
ment shapes its size and structure in Arabidopsis. Plant J. 97,
805–824 (2019).
92. Mboene Noah, A. et al. Dynamics of auxin and cytokinin metabolism
during early root and hypocotyl growth in theobroma cacao. Plants
10, 967 (2021).
93. Cove, D. J. et al. The moss physcomitrella patens: a novel model
system for plant development and genomic studies. Cold Spring
Harb. Protoc. 2009, pdb.emo115 (2009).
94. Vandenbussche,F.,Vaseva,I.,Vissenberg,K.&VanDerStraeten,D.
Ethylene in vegetative development: a tale with a riddle. N. Phytol.
194,895–909 (2012).
95. Bates,D.,Mächler,M.,Bolker,B. & Walker, S. Fitting linear mixed-
effects models using lme4. J. Stat. Softw. 67,1–48 (2015).
96. Kuznetsova, A., Brockhoff, P. B. & Christensen, R. H. B. lmerTest
package: tests in linear mixed effects models. J. Stat. Softw. 82,
1–26 (2017).
97. Lenth,R.V.etal.emmeans:estimatedmarginalmeans,akaleast-
squares means. https://cran.r-project.org/web/packages/
emmeans/emmeans.pdf (2023).
Acknowledgements
We acknowledge Pavel Škaloud (Department of Botany, Charles Uni-
versity, Prague, Czechia) for providing the CAUP algal cultures and
valuable advice; Norbert Zlámal (Botanical Garden, Comenius Uni-
versity, Bratislava, Slovakia) for identifying and providing Selaginella
uncinata; Marie Korecká (Institute of Experimental Botany, Czech Acad-
emy of Sciences, Prague, Czechia) for assistance with experimental
work. This work was supported by Czech Science Foundation project no.
20-13587S, and Charles University Grant Agency (GAUK), project no.
393422. D.V.D.S. acknowledges the Research Foundation Flanders
(FWO; G082421N) and Ghent University (BOF-BAS) for financial support.
Author contributions
V.S. and R.S. cultivated the biological material, analyzed data, and wrote
the text. K.K. and S.H. cultivated and prepared samples of certain ana-
lyzed organisms. R.V. and P.D. performed mass spectrometry. T.D. and
A.P. measured ethylene production. V.S. and S.V. performed the statis-
tical analysis. V.M., D.V.D.S. and J.P. supervised the work and writing.
Competing interests
The authors declare no competing interests.
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