Further characterization and pathogenicity of Didymella microchlamydospora causing stem necrosis of Morus nigra in Iran

Abstract

In the last decade, canker and dieback diseases have caused disease of ornamental and fruit trees of Khuzestan Province in the southwest of Iran. Fourty-eight symptomatic branches and trunks were sampled and a survey was made to identify the probable pathogens, which led to the isolation of the recently established species, Didymella microchlamydospora. A multi-locus DNA sequence based phylogeny, in combination with morphology, was used to characterize seven isolates of this species. Two phylogenetic trees constructed based on the combined sequences of ITS/LSU/tub2 and ITS/LSU/tub2/rpb2 regions showed very little differences, and both trees presented generally consistent relationships among the strongly supported clades. In both of three- and four-locus based phylogenetic trees, our isolates and a reference strain, D. microchlamydospora CBS 105.95, formed supportive monophyletic clades with strong 99% and 100% BS support, respectively. In pathogenicity tests, the isolate of D. microchlamydospora SCUA 14_Dez_Mor formed the necrosis and wood discoloration on stem fragments of Morus nigra. To our knowledge, this is the first report of pathogenicity of D. microchlamydospora on Morus nigra and its association on plants of olive, bitter orange, oleander and bottlebrush worldwide. In addition, we gave a slightly amended description of this species.

Full text

Submitted 10 March 2017, Accepted 11 April 2017, Published 9 June 2017
Corresponding Author: Mehdi Mehrabi-Koushki – e-mail – mhdmhrb@scu.ac.ir; mhdmhrb@gmail.com 835
Further characterization and pathogenicity of Didymella
microchlamydospora causing stem necrosis of Morus nigra in Iran
S. Akram Ahmadpour, Reza Farokhinejad and Mehdi Mehrabi-Koushki
Plant Protection Department, Agriculture Faculty, Shahid Chamran University of Ahvaz, Khuzestan Province, Iran
Ahmadpour SA, Farokhinejad R, Mehrabi-Koushki M 2017 – Further characterization and
pathogenicity of Didymella microchlamydospora causing stem necrosis of Morus nigra in Iran.
Mycosphere 8(7), 835–852, Doi 10.5943/mycosphere/8/7/3.
Abstract
In the last decade, canker and dieback diseases have caused disease of ornamental and fruit
trees of Khuzestan Province in the southwest of Iran. Fourty-eight symptomatic branches and
trunks were sampled and a survey was made to identify the probable pathogens, which led to the
isolation of the recently established species, Didymella microchlamydospora. A multi-locus DNA
sequence based phylogeny, in combination with morphology, was used to characterize seven
isolates of this species. Two phylogenetic trees constructed based on the combined sequences of
ITS/LSU/tub2 and ITS/LSU/tub2/rpb2 regions showed very little differences, and both trees
presented generally consistent relationships among the strongly supported clades. In both of three-
and four-locus based phylogenetic trees, our isolates and a reference strain, D. microchlamydospora
CBS 105.95, formed supportive monophyletic clades with strong 99% and 100% BS support,
respectively. In pathogenicity tests, the isolate of D. microchlamydospora SCUA 14_Dez_Mor
formed the necrosis and wood discoloration on stem fragments of Morus nigra. To our knowledge,
this is the first report of pathogenicity of D. microchlamydospora on Morus nigra and its
association on plants of olive, bitter orange, oleander and bottlebrush worldwide. In addition, we
gave a slightly amended description of this species.
Key words – die back – Khuzestan – multi-locus phylogeny
Introduction
Canker and dieback diseases are common, widespread, and destructive on a wide range of
woody plants (Shurtleff 1997, Horst 2013). These diseases are caused by several fungal taxa
belonging to the different families including Botryosphaeriaceae and Didymellaceae (Phillips et al.
2013, Chen et al. 2015) in Dothideomycetes and Cytospora and Diaporthe in Sordariomycetes
(Sinclair et al. 1987, Lawrence et al. 2015). In the last decade, these diseases have threatened the
ornamental and fruit trees of Khuzestan Province in the southwest of Iran. The potential pathogens
infect all woody plants, especially those low in vigor. The disease causes stem necrosis and canker,
wilting and dieback of twigs and branches (unpublished data).
The species D. microchlamydospora (Aveskamp & Verkley) Q. Chen & L. Cai (formerly
known Phoma microchlamydospora) belongs to the recently established family Didymellaceae (de
Gruyter et al. 2009, Hyde et al. 2013), which includes many taxa previously classified in the genus
Phoma and their related taxa (Chen et al. 2015). This species has been isolated from leaves of
Eucalyptus sp. and an unknown plant (Aveskamp et al. 2009). The genus Didymella sensu lato was
Mycosphere 8(7): 835–852 (2017) www.mycosphere.org ISSN 2077 7019
Article
Doi 10.5943/mycosphere/8/7/3
Copyright © Guizhou Academy of Agricultural Sciences

836
established by Saccardo (1880) to accommodate D. exigua (Niessl) Sacc. (Holm 1975, Corlett
1981). This genus was originally placed in Mycosphaerellaceae, and then subsequently reclassified
in the Pleosporaceae, Phaeosphaeriaceae and Venturiaceae (Hyde et al. 2013, Wijayawardene et
al. 2014, Chen et al. 2015). In recent years, phylogenetic studies have resulted in the dramatic
taxonomic changes in Didymella and other Phoma-like taxa (Aveskamp et al. 2009, 2010, de
Gruyter et al. 2009, Woudenberg et al. 2009, Chen et al. 2015). In order to resolve phylogenetic
relationships and improve the systematics of Phoma and allied genera, ITS, LSU, tub2 and rpb2
sequences were used for species demarcation (Aveskamp et al. 2009, 2010, Woudenberg et al.
2009, Chen et al. 2015). According to the most recent phylogenetic analysis of Phoma-like taxa
(Chen et al. 2015, Hyde et al. 2016), Didymella sensu stricto (Didymella Sacc. ex Sacc., Syll. Fung.
1: 545. 1882. emend. Q. Chen & L. Cai.) was accommodated in the recently introduced family of
Didymellaceae (de Gruyter et al. 2009). The molecular phylogenetic studies showed that the family
Didymellaceae includes most members of Phoma and related asexual genera including the new
emended and introduced genera of Phoma, Ascochyta, Didymella, Epicoccum, Stagonosporopsis,
Allophoma, Heterophoma, Boeremia, Paraboeremia, Macroventuria, Phomatodes, Calophoma,
Leptosphaerulina, Neoascochyta, Xenodidymella, Nothophoma, Neodidymelliopsis, Neodidymella
and Neomicrosphaeropsis. Didymella had first been identified as paraphyletic taxon within the
Didymellaceae (Aveskamp et al. 2010), then a comprehensive phylogenetic analysis of
Didymellaceae was carried out (Chen et al. 2015), and in which Didymella was emended as
monophyletic genus to accommodate 35 known and two unknown species. In the Chen et al. (2015)
study, the genus Didymella was emended to accommodate the species of Didymella exigua, D.
microchlamydospora, D. acetosellae, D. aliena, D. americana, D. anserina, D. arachidicola, D.
aurea, D. bellidis, D. boeremae, D. calidophila, D. chenopodii, D. coffeae-arabicae, D. dactylidis,
D. dimorpha, D. eucalyptica, D. gardeniae, D. heteroderae, D. lethalis, D. longicolla, D.
macrostoma, D. maydis, D. molleriana, D. musae, D. negriana, D. pedeiae, D. pinodes, D.
pomorum, D. rhei, D. viburnicola, D. rumicicola, D. sancta, D. senecionicola, D. subglomerata, D.
subherbarum, D. curtisii, D. glomerata, D. nigricans, D. pinodella, D. protuberans and two
unidentified species. Recently, Didymella cirsii was added (Liu et al. 2015).
The genus Didymella is widely distributed in field and ornamental crops as well as in wild
plants (Chen et al. 2015). The species of this genus are mainly saprobes that are commonly found in
living or dead aerial parts of herbaceous and wooden plants (Chen et al. 2015); some of them also
act as mutualistic endophytes with some plant species (Rayner 1922). Very little is known about the
pathogenicity of Didymella sensu stricto. However, a small number of species belonging to newly
recombined genus of Didymella was reported as plant pathogen (Tivoli and Banniza 2007, Barilli et
al. 2016). The species Didymella pinodes (formerly known Mycosphaerella pinodes) was reported
as main causal agent of Ascochyta blight, one of the most important fungal diseases of pea
worldwide (Tivoli and Banniza 2007, Barilli et al. 2016). In addition, Didymella tanaceti (Syn:
Microsphaeropsis tanaceti haplotype I) and D. rosea (Syn: M. tanaceti haplotype II) were reported
as plant pathogens, that caused tan spot of pyrethrum (Pearce et al. 2016).
According to the current literature (Aveskamp et al. 2009, Chen et al. 2015), two known
strains of Didymella microchlamydospora (CBS 105.95 and CBS 491.90) were regarded as
saprobes (Chen et al. 2015). In this study change it is evident that this species can cause dieback
and necrosis. Here, seven isolates of Didymella microchlamydospora were identified using
phylogenetic analysis based on ITS, LSU, tub2 and rpb2 sequence data. The morphology and
pathogenicity of these isolates is also characterized.
Materials & Methods
Collection of specimens
The specimens were collected from the township of Andimeshk, Ahvaz and Dezful
Khuzestan Province in the southwest of Iran. This climate is hot semi-arid (Koppen climate
classification BSh) with extremely hot summers and mild winters. These areas are generally very

837
hot and occasionally humid, while summertime temperatures routinely exceed 45C and in the
winter, it can rarely drop below freezing. Rainfall is almost exclusively confined to the period from
November to April. During 2015–2016, 48 symptomatic branches and trunks were sampled from
the trees of olive (Olea spp), bitter orange (Citrus aurantium), blackberry (Morus nigra), oleander
(Nerium oleander) and Bottlebrush (Callistemon viminalis), with the symptoms of dieback,
yellowing and defoliation (Fig. 1). The samples were packed in paper bags and transferred to the
lab.
Figure 1 – a The symptoms of die back and decline on Citrus aurantium. b The symptoms of stem
canker and wood discoloration on Morus nigra. c Necrosis and discoloration of branches in
Callistemon viminalis. d Pathogenecity test, necrosis symptom on a stem fragment of Morus nigra
caused by pathogenic isolate of D. microchlamydospora SCUA 14_Dez_Mor (top) compared to a
control fragment (bottom).
Isolation and purification
The small pieces (0.3–1 cm) from healthy and discolored margins of symptomatic branches
and dead stems were excised and surface-sterilized by dipping them in 2% sodium hypochlorite (2–
4 minutes), followed by washing three times with sterile distilled water (2 min). Then, the
fragments were plated on petri plates containing potato dextrose agar (PDA, Difco, USA)
supplemented with streptomycin (30 mg/L). The plates were incubated up to 5–15 days at 28 C,
and individual colonies were cultured to PDA. The isolates were purified by single spore method.
The spore suspension was prepared and 100 μL of which plated on a ¼-strenght PDA. The plates
incubated in the dark at 28 C for 24–48 hours and individual small colonies sub-cultured on PDA as
single-spore isolates. The living cultures of the isolates were deposited in the Collection of Fungal
Cultures, Department of Plant Protection, Shahid Chamran University of Ahvaz, Iran (SCUA 11-
SCUA 17).
Microscopy and growth indicators
The isolates of Didymella were grown on potato dextrose agar (PDA, Merck) and corn meal
agar (CMA, Sigma Aldrich) at 28 C, with 12 hours fluorescent light and 12 hours darkness. The
diameter of colonies was daily measured up to 10-day incubation. Morphological characters were
made at 3–25 days post-inoculation and the colour rate was determined according to the Methuen
handbook of color (Kornerup & Wanscher 1967). The microscopic preparations were made by

838
using the method of Riddle (1950) and Measurements were carried out with the 40× and 100×
objective lens of a Leitz wetzlar (SM-LUX) Basic Biological Light Microscope. The sizes of
characteristic structures were recorded with 50–70 measurements for each structure. The
photomicrographs were made with an OLYMPUS BX51 microscope fitted with an OLYMPUS
DP12 digital camera. Macroscopic and microscopic morphological characters were used to
compare the isolated fungal taxa with the assistance of current mycological literature (Aveskamp et
al. 2009, Chen et al. 2015). Then, for accurate identification, the isolates were subjected to DNA
analysis.
Pathogenicity test
The stem fragments of each trees with similar height, diameter, and vigor were selected. After
surface sterilizing the fragments with 2% sodium hypochlorite (2–4 min) and washing by sterilized
distilled water, a 3-mm-diameter hole was made to the depth of the cambium at 2–3 cm from both
sides of each stem using a scalper. A small quantity of inoculum taken from active-growing edge of
the colonies Didymella microchlamydospora isolates was inoculated into each wound. Free culture
media was placed into wounds as control. The replicates of each treatment were separately placed
into water containing desiccators, sterilized as moist chamber. The desiccators were incubated at
28°C for 3 to 6 weeks after inoculation. Pathogenicity of each isolate were evaluated 3 to 6 weeks
after inoculation by indicating: (i) the presence or absence of callus around the wound, (ii) the
growth and sporulation of fungus in bark surrounding the inoculation point, (iii) the extent of
external longitudinal spread of lesions and (iv) the internal longitudinal spread of discoloration in
xylem.
DNA extraction and amplification
The mycelial biomass of Didymella isolates grown into flasks containing potato dextrose
broth (PDB) was harvested by passing through sterilized filter papers. The mycelia were freeze-
dried (Freeze-Dryer, Alpha 1-2LD Plus, Christ) and powdered in the mortar containing liquid
nitrogen. The genomic DNA was isolated according to modified method established by Reader and
Broda (1985). The mycelial powder was lysed with a lysis buffer and then extracted three times by
Phenol:chloroform:isoamyl alcohol. The genomic DNA was recovered through ethanol-
precipitation typical method. The DNAs were qualified and quantified using Spectrophotometer
(Eppendorf BioPhotometer plus) and loading on the gel. The partial regions of ITS-LSU, rpb2 and
tub2 were amplified using the primer pairs of ITS1/ NL4 (White et al. 1990, O’Donnell 1993),
RPB2-5F2/ fRPB2-7cR (Liu et al. 1999, Sung et al. 2007) and Btub2Fd/ Btub4Rd (Woudenberg et
al. 2009), respectively. PCR reactions were completed in 50 μL final volumes and consisted of 5
μL 10× prime Taq Reaction Buffer (GenBio, South Korea), 6 μL MgCl2 (25mM), 0.6 μL Prime
Taq DNA Polymerase (5U/ μ), 2 μL of each primer (10mM), 2 μL dNTP (10mM mix), 100–500ng
DNA and miliqure water up to 50 μL. The amplification were performed in a thermocycler (MJ
MiniTM Gradient Thermal Cycler) and run with a temperature profile described in the following: the
PCR cycling were for ITS-LSU amplification, initial melting at 94 C for 5 minutes, 35 cycles each
of 30 seconds at 94 C, 30 seconds at 57 C, and 90 seconds at 72 C and followed with a final
extension at 72 C for 10 minutes, for the tub2 amplification, initial melting at 94 C for 5 minutes,
35 cycles each of 30 seconds at 94 C, 30 seconds at 58 C, and 60 seconds at 72 C and followed
with a final extension at 72 C for 10 minutes and for the rpb2 amplification, initial melting at 94 C
for 5 minutes, 35 cycles each of 30 seconds at 94 C, 30 seconds at 57 C, and 60 seconds at 72 C
and followed with a final extension at 72 C for 10 minutes.
Sequencing and phylogenetic analyses
PCR products were purified through ethanol-precipitation method (Crouse & Amorese 1987)
and then sequenced using forward and reverse primers by Macrogen Company. The Sequences
obtained from each primer pairs were assembled using DNA Baser Sequence Assembeler v4
programs (2013, Heracle BioSoft, www.DnaBaser.com). The phylogenetic analysis of Didymella

839
microchlamydospora isolates was carried out with including the reference sequences belonging to
the known genera of Didymellaceae and species of Didymella (225 available sequences mostly
from Aveskamp et al. 2010 and Chen et al. 2015 included) (Table 1). The species of Pleospora
betae were used as outgroup taxon to root phylogenetic trees.
The sequences of ITS, LSU, tub2 and rpb2 were aligned individually using ClustalW in
BioEdit v. 7.0.9.0 (Hall 1999), trimmed to the same starting position and then assembled. The
combined ITS-LSU-tub2 and ITS-LSU-tub2-rpb2 datasets were multiple-aligned using ClustalW in
BioEdit v. 7.0.9.0 (Hall 1999). Phylogenetic analysis was performed with maximum parsimony and
maximum likelihood algorithm. Phylogenetic trees were constructed using MEGA version 6
(Tamura et al. 2013). Best-fitting ML nucleotide substitution model for each dataset was
determined using the model test function in MEGA version 6. The phylogenetic trees were
constructed with Subtree-Pruning-Regrafting (SPR) algorithm and following options: Gaps
(insertion/deletions) were treated as missing data, Bootstrap (BP) analyses were done with 1000
replicates, Initial Trees for ML were made by NJ/BioNJ algorithm and Branch Swap Filter was set
very strong. Two final alignments used for phylogenetic analyses were deposited in TreeBASE
(http://purl.org/phylo/treebase/phylows/study/TB2:S21100).
Results
Morphological characterization (Fig. 2)
Hyphae diameter in 14-day colonies 2.5–4 μm ( = 3.2 μm, n = 50). Conidiomata pycnidial,
pycnidia mostly solitary or aggregated, superficial on or submerged into the agar, dark brown, with
age becoming darker, variable in shape and size (macro-and micro-pycnidium). Macropycnidia
globose, glabrous or covered with hyphal outgrows, 100–190 × 100–190 μm ( = 139 × 139 μm, n
= 50) (Fig. 2). Ostioles 1–3, papillate, rarely on a distinct neck. Pycnidial wall
pseudoparenchymatous, composed of oblong to isodiametric cells, 2–5 layers. Micropycnidia
globose to subglobose, glabrous or covered with hyphal outgrows, 50–80 × (40–)49–70(–80) μm (
= 61 × 59 μm, n = 50). Conidia hyaline to pale brown, smooth- and thin-walled, subglobose to
ellipsoidal, aseptate and guttulate, (2.5)3–5.5(6) × (1.5)2–3.2(3.8) μm ( = 4.3 × 2.4 μm, n = 70).
Chlamydospores mostly unicellular, solitary or in chain, intercalary or terminal, smooth, brown,
globose to subglobose, (3)4–7.5(10) × (2.5)3–7.5 μm ( = 5.9 × 4.6 μm, n = 50). Multicellular
Chlamydospores (pseudosclerotioid and dictyosporous) variable in shape and size, brown,
intercalary, sparse and solitary, smooth.
Colonies on PDA, 70–80 mm diameter after 10 days of incubation at 28±0.5 C, blackish-
brown with whitish cream margins at early growth stage, with age becoming blackish green in the
central and olivaceous green in the edge, staining the agar in pink collar due to the production of a
diffusible pigment, floccose growth, the rings of sporulation containing black pycnidia becoming
darker and compacter towards the center of the colony; reverse blackish green with creamy to
orange edges, leaden black in zones with abundant pycnidia, darkening towards the center of the
colony. Colonies on CMA, 65–75 mm diameter after 10 days of incubation at 28 ± 0.5 C, grey to
brownish grey with lighter edge, smooth, the pycnidia appear as scattered small dots of brown to
black or rings of sporulation; reverse grey to olivaceous green with lighter edge, leaden blackish
brown in pycnidia containing zone.
Material examined – IRAN, Khuzestan Province, Andimeshk, on dead branch of Olea
europaea, 11 August 2015, S.A. Ahmadpour (SCUA 11_And_ Ole); Ahvaz, on dead branch of
Olea europaea, 12 Oceober 2015, S.A. Ahmadpour (SCUA 12_Ahw_Ole); on dead branch of
Citrus aurantium, 12 Oceober 2015, S.A. Ahmadpour (SCUA 13_Ahw_Cit); on dead branch of
Olea sp, 12 Oceober 2015, S.A. Ahmadpour (SCUA 16_Ahv_Ole); on dead branch of Callistemon
viminalis, 12 Oceober 2015, S.A. Ahmadpour (SCUA 15_Ahv_Cal); on dead branch of Nerium sp,
12 Oceober 2015, S.A. Ahmadpour (SCUA 12_Ahv_Ner); Dezful, on dead branch of Morus nigra,
15 August 2015, S.A. Ahmadpour (SCUA 14_Dez_Mor).

840
Table 1 Strains used in this study and their GenBank accession numbers. Newly generated sequences are indicated in bold.
Species name
Isolate name or
strain no.
Source
Origin
GenBank Accession number
ITS
LSU
rpb2
tub2
Didymella
microchlamydospora
IRAN 2788C;
SCUA 11_And_Ole
Olea europaea
Iran
KX139019
KX139028
KY464923
KY449026
D. microchlamydospora
IRAN 2789C;
SCUA 12_Ahw_Ole
Olea europaea
Iran
KX139018
KX139027
KX821250
KX821247
D. microchlamydospora
IRAN 2790C;
SCUA 13_Ahw_Cit
Citrus aurantium
Iran
KX139014
KX139023
KX821249
KX821246
D. microchlamydospora
SCUA 14_Dez_Mor
Morus nigra
Iran
KX139012
KX139021
KX821248
KX821245
D. microchlamydospora
IRAN 2791C;
SCUA 16_Ahv_Ole
Olea sp.
Iran
KY449004
KY449013
-
-
D. microchlamydospora
SCUA 15_Ahv_Cal
Callistemon viminalis
Iran
KY449005
KY449014
-
-
D. microchlamydospora
IRAN 2792C;
SCUA 12_Ahv_Ner
Nerium sp.
Iran
KY449006
KY449015
-
-
D. exigua
CBS 183.55
Rumex arifolius
France
GU237794
EU754155
EU874850
GU237525
D. acetosellae
CBS 179.97
Rumex hydrolapathum
The Netherlands
GU237793
GU238034
KP330415
GU237575
D. aliena
CBS 379.93
Berberis sp.
The Netherlands
GU237851
GU238037
KP330416
GU237578
D. americana
CBS 185.85
Zea mays
USA
FJ426972
GU237990
KT389594
FJ427088
D. anserina
CBS 253.80
-
Germany
KT389498
KT389715
KT389595
KT389795
D. arachidicola
CBS 333.75
Arachis hypogaea
South Africa
GU237833
GU237996
KT389598
GU237554
D. aurea
CBS 269.93
Medicago polymorpha
New Zealand
GU237818
GU237999
KT389599
GU237557
D. bellidis
CBS 714.85
Bellis perennis
The Netherlands
GU237904
GU238046
KP330417
GU237586
D. boeremae
CBS 109942
Medicago littoralis
Australia
FJ426982
GU238048
KT389600
FJ427097
D. chenopodii
CBS 128.93
Chenopodium quinoa
Peru
FJ427060
GU238053
-
GU237591
D. coffeae-arabicae
CBS 123380
Coffea arabica
Ethiopia
FJ426993
GU238005
KT389603
FJ427104
D. curtisii
PD 92/1460
Sprekelia sp.
The Netherlands
FJ427041
GU238012
KT389604
FJ427151
D. eucalyptica
CBS 377.91
Eucalyptus sp.
Australia
GU237846
GU238007
KT389605
GU237562
D. exigua
CBS 183.55
Rumex arifolius
France
GU237794
EU754155
EU874850
GU237525
D. microchlamydospora
CBS 105.95
Eucalyptus sp.
UK
FJ427028
GU238104
KP330424
FJ427138
D. rhei
CBS 109177
Rheum rhaponticum
New Zealand
GU237743
GU238139
KP330428
GU237653
D. rumicicola
CBS 683.79
Rumex obtusifolius
New Zealand
KT389503
KT389721
KT389622
KT389800
D. sancta
CBS 281.83
Ailanthus altissima
South Africa
FJ427063
GU238030
KT389623
FJ427170

841
Table 1 (continued)
Species name
Isolate name
or strain no.
Source
Origin
GenBank Accession number
ITS
LSU
rpb2
tub2
Didymella sp. 1
CBS 379.96
Pteris sp.
The Netherlands
KT389504
KT389722
KT389624
KT389801
Didymella sp. 2
CBS 115.58
Chrysanthemum roseum
Germany
KT389505
KT389723
KT389625
KT389802
D. subglomerata
CBS 110.92
Triticum sp.
USA
FJ427080
GU238032
KT389626
FJ427186
D. viburnicola
CBS 523.73
Viburnum cassioides
The Netherlands
GU237879
GU238155
KP330430
GU237667
D. negriana
CBS 358.71
Vitis vinifera
Germany
GU237838
GU238116
KT389610
GU237635
D. nigricans
PD 77/919
Actinidea chinensis
New Zealand
GU237915
GU238001
KT389611
GU237559
D. pedeiae
CBS 124517
Schefflera elegantissima
The Netherlands
GU237770
GU238127
KT389612
GU237642
D. pinodella
CBS 531.66
Trifolium pretense
USA
FJ427052
GU238017
KT389613
FJ427162
D. pinodes
CBS 525.77
Pisum sativum
Belgium
GU237883
GU238023
KT389614
GU237572
D. protuberans
CBS 377.93
Daucus carota
The Netherlands
GU237847
GU238014
KT389619
GU237565
D. molleriana
CBS 229.79
Digitalis purpurea
New Zealand
GU237802
GU238067
KP330418
GU237605
D. exigua
CBS 183.55
Rumex arifolius
France
GU237794
EU754155
EU874850
GU237525
D. lethalis
CBS 103.25
-
-
GU237729
GU238010
KT389607
GU237564
D. mascrostoma
CBS 482.95
Larix decidua
Germany
GU237869
GU238099
KT389609
GU237626
D. maydis
CBS 588.69
Zea mays
USA
FJ427086
EU754192
GU371782
FJ427190
D. calidophila
CBS 448.83
Soil
Egypt
FJ427059
GU238052
-
FJ427097
D. dactylidis
CBS 124513
Dactylis glomerata
USA
GU237766
GU238061
-
GU237599
D. dimorpha
CBS 346.82
Opuntiae sp
Spain
GU237835
GU238068
-
GU237606
D. gardeniae
CBS 626.68
Gardenia jasminoides
India
FJ427003
GQ387595
KT389606
FJ427114
D. glomerata
CBS 528.66
Chrysanthemum sp.
The Netherlands
FJ427013
EU754184
FJ427013
FJ427124
D. heteroderae
CBS 109.92
Undefined material
The Netherlands
FJ426983
GU238002
KT389601
FJ427098
Neodidymelliopsis cannabis
CBS 234.37
Cnnabis sativa
-
GU237804
GU237961
KP330403
GU237523
Xenodidymella applanata
CBS 205.63
Rubus idaeus
The Netherlands
GU237798
GU237998
KP330402
GU237556
Paraboeremia adianticola
CBS 187.83
Polystichum adiantiforme
USA
GU237796
GU238035
KP330401
GU237576
Ascochyta pisi
CBS 122751
Pisum sativum
Canada
KP330432
KP330444
EU874867
KP330388
Phomatodes aubrietiae
CBS 627.97
Aubrietia sp.
The Netherlands
GU237895
GU238045
KT389665
GU237585
Calophoma clematidina
CBS 102.66
Clematis sp.
UK
FJ426988
FJ515630
KT389587
FJ427099
Phoma herbarum
CBS 377.92
Human leg
The Netherlands
KT389536
KT389756
KT389663
KT389837
Macroventuria anomochaeta
CBS 525.71
Decayed canvas
South Africa
GU237881
GU237984
GU456346
GU237544
Leptosphaerulina australis
CBS 317.83
Eugenia aromatica
Indonesia
GU237829
EU754166
GU371790
GU237540

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Table 1 (continued)
Species name
Isolate name or
strain no.
Source
Origin
GenBank Accession number
ITS
LSU
rpb2
tub2
Epicoccum nigrum
CBS 125.82
Human toenail
The Netherlands
FJ426995
GU237974
KT389631
FJ427106
Stagonosporopsis hortensis
CBS 104.42
-
The Netherlands
GU237730
GU238198
KT389680
GU237703
Allophoma tropica
CBS 436.75
Saintpaulia ionantha
Germany
GU237864
GU238149
KT389556
GU237663
Heterophoma adonidis
CBS 114309
Adonis vernalis
Sweden
KT389506
KT389724
KT389637
KT389803
Neoascochyta exitialis
CBS 118.40
-
-
KT389514
KT389732
KT389647
KT389812
Pleospora betae
CBS 523.66
Beta vulgaris
The Netherlands
FJ426981
EU754179
KT389670
KT389842
1 Abbreviation of culture collections: CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; PD: Plant Protection Service,
Wageningen, the Netherlands; IRAN: Iranian Fungal Culture Collection, Iranian Research Institute of Plant Protection, Iran; SCUA: the Collection of
Fungal Cultures, Department of Plant Protection, Shahid Chamran University of Ahvaz, Iran.
DNA analysis and phylogenetic characterization
The sequences of ITS, LSU, tub2 and rpb2 belonging to the isolates under study were submitted to GenBank (table 1) under the generic name
Didymella microchlamydospora. These isolates shared 98.8% sequence identity in the ITS region (430 bp) attributed to 2 SNPs and three bp
insertion/deletion, 100% sequence identity in the LSU region (590 bp), 99.7% sequence identity in the tub2 region (306 bp) attributed to one SNPs,
and 99 % sequence identity in the rpb2 region (782 bp) attributed to eight SNPs. Using a BLASTn search, the ITS sequences of seven D.
microchlamydospora isolates showed 99–100% sequence identity to reference strain D. microchlamydospora CBS 105.95.
Sixty-two and 55 taxa, including all described species of Didymella and a type species from all the known genera of Didymellaceae, were
included in the three-locus and four-locus based phylogeny, respectively (Table 1). The composite sequence alignment was 1206 and 1809 characters
in length, including alignment gaps (ITS: 420 bp, LSU: 500 bp, tub2: 286 bp, rpb2: 603 bp) for three and four regions, respectively. Of those
characters 1233 bp (ITS: 324 bp, LSU: 450 bp, tub2: 176 bp, rpb2: 283 bp) were constant and 576 bp (ITS: 96 bp, LSU: 50 bp, tub2: 110 bp, rpb2: 320
bp) were variable. The best-fitting ML nucleotide substitution model for phylogenetic analysis of three-locus and four-locus combined datasets were
selected Tamura-Nei (TN93+G+I) and General Time Reversible (GTR+G+I) models, respectively. The phylogenetic trees of the maximum likelihood
analysis based on both combined datasets are shown in Figs 3 and 4. The topology of phylogenetic trees showed very little differences, and both trees
presented generally consistent relationships among the strongly supported clades (Figs 3, 4). The trees topology of both three- and four-locus
phylogenetic analysis provided the evidence that the isolates under study were associated with the species Didymella microchlamydospora. In both
trees, our isolates and a reference strain from GenBank, D. microchlamydospora CBS 105.95, generated supportive monophyletic clades with strong
BS 99% and 100% support. In both trees, the reference strain of Neoascochyta exitialis CBS 118.40 among the representative members of the family
Didymellaceae positioned as a basal taxon. In addition, the trees obtained through maximum parsimony analysis supported the tree obtained from ML
analysis (not shown).

843
Figure 2 – Didymella microchlamydospora isolate SCUA 14_Dez_Mor. a, b Colony on PDA
(front and reverse). c, d Colony on CMA (front and reverse). e, f, g, h Pycnidia formed on PDA and
CMA. i, j, k, l Chlamydospores. m Conidia.
Ecology and distribution
In the last decade, some of the decline symptoms including; yellowing, wilting, defoliation,
dieback and canker were observed on various tree species in whole area of investigation. This
disease affected about 5% of the various ornamental and fruit trees such as Citrus spp., Eucalyptus
spp., Morus spp., Conocarpus erectus, Ziziphus nummularia, Nerium oleander, Juglans regia,
Prosopis spicigera, Cupressus semperviren, Punica grenatum, Prosopis stephaniana, Olea
europaea, Callistemon viminalis, Bauhinina purpurea, Albizia lebbeck and Cordia mixa. The first
observed signs in affected trees were dieback, and in which the dead of infected tissues resulted in
the girdling of shoots and branches. Following, the causal fungus developed internally and
destroyed the growth rings, which is the characteristic of other stem canker causing agents. Death
of branches throughout the crown led to gradual tree decline or the tree was dying starting from the
top of the crown. An attempt was made to identify the potential canker pathogens and other
associated agents. In all, 48 samples were surveyed, 49 isolates of potential pathogenic fungi were
detected (unduplicated data), and seven isolates were identified as Didymella microchlamydospora.
The isolates of Didymella microchlamydospora SCUA 11-And_Ole and SCUA 12-Ahv_Ole were

844
Figure 3 – Phylogenetic tree constructed from a maximum likelihood analysis based on a
concatenated alignment of ITS, LSU and tub2 sequences of four Didymella microchlamydospora
isolates under study and 16 type strains representing a type species of each described genus of
Didymellaceae and 42 described species of genus Didymella downloaded from GenBank. Bootstrap
values greater than 50% (expressed as percentages of 1000 replications) are shown at the nodes.
The tree was rooted with Pleospora betae CBS 523.66.

845
Figure 3 (continued)

846
Figure 4 – Phylogenetic tree constructed from a maximum likelihood analysis based on a
concatenated alignment of ITS, LSU, tub2 and rpb2 sequences of four Didymella
microchlamydospora isolates under study and 16 type strains representing a type species of each
described genus of Didymellaceae and 35 described species of genus Didymella downloaded from
GenBank. Bootstrap values greater than 50% (expressed as percentages of 1000 replications) are
shown at the nodes. The tree was rooted with Pleospora betae CBS 523.66.

847
Figure 4 (continued)

848
firstly isolated from dead branches of olive (Olea europaea, Oleaceae) in Andimeshk and
Ahvaz, and then subsequently, D. microchlamydospora SCUA 13-Ahv_Cit from Bitter orange
(Citrus aurantium, Rutaceae) in Ahvaz, D. microchlamydospora SCUA 14-Dez_Mor from
blackberry (Morus nigra, Moraceae) in Dezful, D. microchlamydospora SCUA 15- Ahv_Ner from
oleander (Nerium oleander, Apocynaceae) in Ahvaz and D. microchlamydospora SCUA 16-
Ahv_Cal from weeping bottlebrush (Callistemon viminalis, Myrtaceae) in Ahvaz.
Pathogenicity tests
Both isolates of Didymella microchlamydospora SCUA 11_And_Ole and D.
microchlamydospora SCUA 14_Dez_Mor were able to grow and sporulate in the bark surrounding
the inoculation point on stem fragments of Olea europaea and Morus nigra, respectively. The
isolate of D. microchlamydospora SCUA 14_Dez_Mor developed the external longitudinal lesion
on the inoculation point three weeks after inoculation, which was associated with wood necrosis
and discoloration in xylem (Fig. 1), while the isolate of D. microchlamydospora SCUA
11_And_Ole did not. In both test plants, the callus was not formed around the inoculation wound.
This pathogenic fungus was re-isolated from necrosis-like areas formed on stem fragments of M.
nigra, and the identity as D. microchlamydospora species was confirmed by morphological
characterization.
Discussion
In our study, seven Didymella microchlamydospora isolates were recovered from 48 plant
species. This is the first report of D. microchlamydospora in Iran. Phoma microchlamydospora
Aveskamp & Verkley, was described by Aveskamp et al. (2009), and then, recombined into
Didymella microchlamydospora by Chen et al. (2015). Here, further morphological and molecular
characterization, pathogenicity on Morus nigra, and a phylogenetic analysis between the isolates
under study and other species within the Didymellaceae was evaluated.
In morphology, our isolates are slightly different from reference strain of D.
microchlamydospora CBS 105.95. The diameter of macropycnidia was less than to the strain D.
microchlamydospora CBS 105.95 (100–190 vs. 150–260 μm) (Aveskamp et al. 2009). In similar to
the reference strain (Aveskamp et al. 2009), our isolates produced ostiolate and papillate pycnidia,
but rarely on a distinct neck as described for D. microchlamydospora CBS 105.95. The width and
length of conidia and unicellular chlamydospores are somewhat different but it cannot be used to
distinguish the species from each other. The numerous measurements in this study and previous
observations (McPartland 1994, Chen et al. 2015) demonstrated that, in general, the conidial length
is much more variable, and the conidial size mostly depends on the location of pycnidia.
Furthermore, Conidia in pycnidia produced on culture have been usually observed somewhat larger
than those from living host (McPartland 1994).
In the current study, the identification of Didymella microchlamydospora isolates based on
morphological characterization and BLAST search algorithm is strongly supported in multi-locus
phylogeny based on the combined regions of ITS, LSU, tub2 and rpb2. Four isolates of D.
microchlamydospora were used in the phylogenetic analyses for constructing two phylograms
based on three-locus (ITS-LSU-tub2) and four-locus (ITS-LSU-tub2-rpb2) based combined
datasets. In both three- and four- locus based phylogenetic trees, sequence dataset worked well to
distinguish closely related species in Didymella and our isolates clustered with reference strain D.
microchlamydospora CBS 105.95, distinct from the other Didymella species (Figs 3, 4). Analysis
of congruence between the ITS, LSU, tub2 and rpb2 loci used in the phylogenetic analysis showed
that the LSU region had the lowest correlation scores with 10% sequence diversity and rpb2 region
had the highest correlation scores with 53% sequence diversity. This was most probably due to the
low resolution provided by the LSU, which was expected due to the nature of its evolution within
species. The LSU locus shared 90% sequence identify between the species of Didymellaceae,
indicating their close phylogenetic relationship. However, the LSU locus of filamentous fungi is
often not sufficient to delimit taxa at the species level (Lumbsch et al. 2000, Eberhardt 2010). Due

849
to the abundant homoplasy in phenotypic characteristics and difficulties in the morphological
identification, it is difficult to distinguish Phoma-like taxa including, the species of Didymella
(Chen et al. 2015). Genealogical concordance analysis using several unlinked DNA loci have been
already resulted in the dramatic taxonomic changes in Phoma and Phoma-like genera (de Gruyter
et al. 2009, 2010, 2012, Aveskamp et al. 2010, Ariyawansa et al. 2015, Chen et al. 2015, Liu et al.
2015, Hyde et al. 2016, Li et al. 2016, Tibpromma et al. 2017) as well as other fungi such as in
Alternaria (Woudenberg et al. 2013), Bipolaris (Manamgoda et al. 2011, 2012), Colletotrichum
(Cannon et al. 2012, Jayawardena et al. 2016), Fusarium (Short et al. 2013, Laurence et al. 2014),
Phyllosticta (Wikee et al. 2011, Hyde et al. 2014), Trichoderma (Druzhinina et al. 2010) and other
taxa in Kingdom Myceteae (Ariyawansa et al. 2015, Crous et al. 2015, Liu et al. 2015, Hyde et al.
2016, Li et al. 2016, Tibpromma et al. 2017). Chen et al. (2015) have indicated, that the combined
sequence of ITS, LSU, tub2 and rpb2 work well in demarcating Didymella species. The results of
these phylogenetic analyses validate the species delimitation of our isolates as D.
microchlamydospora.
In pathogenicity tests, of the four tested isolates, D. microchlamydospora SCUA
14_Dez_Mor formed the necrosis symptom on stem fragments of Morus nigra (Fig. 1). Dark brown
to black discoloration expanded rapidly in a longitudinal direction. Previous studies have shown
that Didymella pinodes on Pisum sativum (Tivoli & Banniza 2007, Barilli et al. 2016) and
Didymella tanaceti and Didymella rosea on pyrethrum plant (Pearce et al. 2016) acts as a
phytopathogen in the UK and Australia, respectively. Chen et al. (2015) and Pearce et al. (2016)
supported the placement of these phytopathogenic species in Didymella sensu stricto. In our study,
one isolate of Didymella microchlamydospora infected plant hosts and developed necrosis
symptoms. To the best of our knowledge, this is the first phyto-pathogenicity report for Didymella
microchlamydospora worldwide.
Observational assessment of areas sampled showed, the disease index and tree mortality
positively correlates with environmental stress. Since drought and extremely hot summers became
more common in Khuzestan during the last decade, higher than usual incidence of die back diseases
may be due to drought stresses and higher annual temperatures that made trees more susceptible to
the disease. Observational assessment showed that there was a clear increase in decline symptoms
in the zones with low fertility soils, deficiency of water, prolonged exposure to extremely high
temperatures, summer sunscald, nutritional imbalances, soil compaction, changes in the soil grade
and mechanical injuries. Previous studies showed that environmental stress, such as high
temperatures and drought periods could play a role in increasing the virulence and expansion of the
Didymellaceae, Botryosphaeriaceae and other decline pathogens (Smith et al. 1996, Kim et al.
2001, Arnold & Herre 2003, Desprez-Loustau et al. 2007, Slippers & Wingfield 2007, Botella et al.
2010, Dissanayake et al. 2015, Fan et al. 2016, Anonym 2017, Delgado-Cerrone 2017).
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