REVIEWpublished: 20 March 2015
doi: 10.3389/fpls.2015.00146
Frontiers in Plant Science | www.frontiersin.org 1 March 2015 | Volume 6 | Article 146
Edited by:
Brigitte Mauch-Mani,
Université de Neuchâtel, Switzerland
Reviewed by:
Sebastian Bartels,
University of Basel, Switzerland
Prashant Singh,
Lancaster University, UK
*Correspondence:
Renato D’Ovidio,
Dipartimento di Scienze e Tecnologie
per l’Agricoltura, le Foreste, la Natura
e l’Energia, Università Degli Studi Della
Tuscia, 01100 Viterbo, Italy
Giulia De Lorenzo,
Dipartimento di Biologia e
Biotecnologie “Charles Darwin,”
Sapienza Università di Roma, Roma,
Italy
Specialty section:
This article was submitted to
Plant-Microbe Interaction, a section of
the journal Frontiers in Plant Science
Received: 05 January 2015
Accepted: 23 February 2015
Published: 20 March 2015
Citation:
Kalunke RM, Tundo S, Benedetti M,
Cervone F, De Lorenzo G and
D’Ovidio R (2015) An update on
polygalacturonase-inhibiting protein
(PGIP), a leucine-rich repeat protein
that protects crop plants against
pathogens. Front. Plant Sci. 6:146.
doi: 10.3389/fpls.2015.00146
An update on polygalacturonase-inhibiting protein (PGIP), aleucine-rich repeat protein thatprotects crop plants againstpathogens
Raviraj M. Kalunke 1, Silvio Tundo 1, Manuel Benedetti 2, Felice Cervone 2,
Giulia De Lorenzo 2* and Renato D’Ovidio 1*
1Dipartimento di Scienze e Tecnologie per l’Agricoltura, le Foreste, la Natura e l’Energia, Università della Tuscia, Viterbo, Italy,2Dipartimento di Biologia e Biotecnologie “Charles Darwin”, Sapienza Università di Roma, Roma, Italy
Polygalacturonase inhibiting proteins (PGIPs) are cell wall proteins that inhibit the
pectin-depolymerizing activity of polygalacturonases secreted by microbial pathogens
and insects. These ubiquitous inhibitors have a leucine-rich repeat structure that is
strongly conserved in monocot and dicot plants. Previous reviews have summarized the
importance of PGIP in plant defense and the structural basis of PG-PGIP interaction;
here we update the current knowledge about PGIPs with the recent findings on the
composition and evolution of pgip gene families, with a special emphasis on legume and
cereal crops. We also update the information about the inhibition properties of single
pgip gene products against microbial PGs and the results, including field tests, showing
the capacity of PGIP to protect crop plants against fungal, oomycetes and bacterial
pathogens.
Keywords: polygalacturonase inhibiting proteins (PGIPs), gene family, transgenic plants, plant protection, fungal
pathogens, bacterial pathogens
Introduction
Successful colonization of plant tissues by microbial pathogens requires the overcoming of the cellwall. To this end, pathogens produce a wide array of plant cell wall degrading enzymes (CWDEs),among which endo-polygalacturonases (PGs; EC 3.2.1.15) are secreted at very early stages of theinfection process (tenHave et al., 1998). PGs cleave the α-(1–4) linkages between theD-galacturonicacid residues of homogalacturonan, the main component of pectin, causing cell separation andmaceration of the host tissue. To counteract the activity of PGs, plants deploy the cell wall poly-galacturonase inhibiting proteins (PGIPs) that inhibit the pectin-depolymerizing activity of PGs.No plant species or mutants totally lacking PGIP activity have been characterized so far. The struc-ture of PGIPs is typically formed by 10 imperfect leucine-rich repeats (LRRs) of 24 residues each,which are organized to form two β-sheets, one of which (sheet B1) occupies the concave innerside of the molecule and contains residues crucial for the interaction with PGs (Di Matteo et al.,2003). In addition to PG inhibition, the interaction between PGs and PGIPs promotes the forma-tion of oligogalacturonides (OGs), which are elicitors of a variety of defense responses (Cervoneet al., 1989; Ridley et al., 2001; Ferrari et al., 2013). Since many aspects of the PGIP biology have
Kalunke et al. PGIP and crop protection
been already summarized in previous reviews (De Lorenzo et al.,2001; De Lorenzo and Ferrari, 2002; D’Ovidio et al., 2004a;Gomathi and Gnanamanickam, 2004; Shanmugam, 2005; DiMatteo et al., 2006; Federici et al., 2006; Cantu et al., 2008;Misas-Villamil and van der Hoorn, 2008; Protsenko et al., 2008;Reignault et al., 2008; Lagaert et al., 2009), here we present anoverview of the recent findings on genome composition and evo-lution of pgip gene families and on the efficacy of PGIP to limitthe development of diseases caused by microbial pathogens incrop plants.
PGIP Genes and their GenomicOrganization
Early characterization of a polygalacturonase-inhibiting activitywas reported in 1970s (Albersheim and Anderson, 1971) andthe first pgip gene was isolated 20 years later in French bean(Toubart et al., 1992). Since then, several PGIPs and a largenumber of pgip genes have been characterized. Up to now morethan 170 complete or partial pgip genes from dicot and mono-cot plants have been deposited in nucleotide databases (e.g.,http://www.ncbi.nlm.nih.gov/). Most of these genes have beenidentified as pgip genes on the basis of sequence identity butonly a few of them have been shown to encode proteins withPG-inhibitory activity.
Genome analysis has shown that pgip genes did not undergoa large expansion and may exist as single genes, as in diploidwheat species (Di Giovanni et al., 2008), or organized into genefamilies, the members of which are organized in tandem andcan vary from two, as in Arabidopsis thaliana (Ferrari et al.,2003), to sixteen, as in Brassica napus (Hegedus et al., 2008).The majority of pgip genes are intronless, however, some ofthem can contain a short intron as in Atpgip1 and Atpgip2 (Fer-rari et al., 2003). Moreover, pgip genes can be inactivated bytransposon elements as in cultivated and wild wheat where theoccurrence of Copia-retrotransposon and Vacuna transposonshas been reported (Di Giovanni et al., 2008). Characterized pgiploci are shown in Figure 1. Like other families of defense-relatedgenes, pgip families show variation in the expression pattern ofthe different members, some of which are constitutive, others aretissue-specific and, in most cases, up-regulated following stressstimuli (see reviews indicated above; Table 1). At the proteinlevel, members of a pgip family show both functional redundancyand sub-functionalization (De Lorenzo et al., 2001; Federici et al.,2006). As suggested previously, these features likely have an adap-tive significance for combating more efficiently a broad array ofpathogens (Ferrari et al., 2003) or responding more rapidly todiverse environmental stimuli (D’Ovidio et al., 2004b). In supportof this view, a recent analysis of the genomic organization andcomposition of the legume pgip families suggested that the forcesdriving the evolution of the pgip genes follow the birth-and-deathmodel (Kalunke et al., 2014), similarly to what proposed for theevolution of NBS-LRR-type R genes (Michelmore and Meyers,1998). This possibility is based on genomic features that includeinferred recent duplications, diversification as well as pseudoge-nization of pgip copies, as found in soybean, bean, barrel cloverand chickpea (Kalunke et al., 2014). The organization of the pgip
families therefore supports the view that tandem duplications arefrequent in stress-related genes and are beneficial for survival inchallenging environments (Oh et al., 2012).
Inhibition Activity of PGIPs
A number of papers deals with the inhibition activity of PGIPspurified from several plant tissues. This aspect has been reviewedseveral years ago (De Lorenzo et al., 2001); here, we presentan update of this information (Table 2). Because purified PGIPsmay contain a mix of highly similar PGIP isoforms, the activ-ity detected in a tissue may result from the contribution of theactivities of different PGIPs expressed in that tissue. An appropri-ate approach to study the inhibition activity of individual PGIPisoforms is their expression in a heterologous system. However,only a few of the more than 170 pgip genes isolated so far fromdifferent plant species have been investigated. As reported inTable 3, individual heterologous expression and analysis of allmembers of a pgip family has been performed only for Ara-bidopsis (Ferrari et al., 2003), common bean (D’Ovidio et al.,2004b), soybean (D’Ovidio et al., 2006; Kalunke et al., 2014)and wheat (Janni et al., 2013). PGIPs have been expressed inprokaryotic systems, as a fusion with the maltose-binding pro-tein (MBP) (Jang et al., 2003; Table 3) or using lower temperaturefor bacterial growth (Chen et al., 2011), in Pichia pastoris and inplants by stable transformation or, transiently, by virus-mediatedexpression (Table 3). In some cases, the proteins were success-fully expressed, but did not show any inhibitory activity in vitro,as, for example, in the case of some GmPGIPs (D’Ovidio et al.,2006). GmPGIP3, but not GmPGIP1, GmPGIP2, and GmPGIP7showed inhibitory activity, whereas no expression of GmPGIP5was obtained (D’Ovidio et al., 2006; Kalunke et al., 2014). Sim-ilarly, TaPGIP1 and TaPGIP2, encoded by the two members ofthe wheat pgip family, were successfully expressed but showed noinhibition activity (Janni et al., 2013).
The absence of inhibition activity in vitro may also reflectthe possibility that some PGIPs are active only in the in plantaenvironment, as suggested by Joubert et al. (2006) in the caseof the Botrytis cinerea BcPG2 and VvPGIP1 from grapevine(Vitis vinifera L.). These proteins do not interact in vitro,although VvPGIP1 reduces symptoms caused by BcPG2 uponco-infiltration in leaves. The number and sources of PGs testedis also limited; only a few studies have been carried out againstPGs of bacteria and insects (Doostdar et al., 1997; D’Ovidio et al.,2004b; Frati et al., 2006; Hwang et al., 2010; Schacht et al., 2011;Kirsch et al., 2012). The limitations of data prevents to draw con-clusions about correlations between PGIPs of specific plant fam-ilies and specific pathogens. Notably, PG produced by a highlydetrimental pathogen, Fusarium verticillioides, is not inhibitedby any known PGIP (see Table 2). This PG has been a target ofan unsuccessful attempt to render PvPGIP2 an efficient inhibitoragainst this PG (see below, Benedetti et al., 2011a).
The utilization of pgip genes for crop protection relies on theidentification of inhibitors with broad specificities against themany PGs produced by phytopathogens and/or the construc-tion of novel PGIPs with stronger and broader inhibitor activity.Many more PGIPs than those reported in Tables 2, 3 exist in
Frontiers in Plant Science | www.frontiersin.org 2 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
FIGURE 1 | Schematic representation of the genomic organization
pgip families in rice, wheat, bean, soybean, chickpea, barrel clover
and thale cress. Each block-arrow with compound-type lines represents a
predicted pgip gene and a block-arrow with dash type lines represents a
predicted pseudo-gene or remnant gene. Vertical line within block-arrow
indicates introns (Capgip2, Atpgip1, and Atpgip2) or a Copia retrotransposon
(Tapgip3). The direction of the arrow indicates ATG to stop codon. The
location of pgip genes of legume species are based on Kalunke et al. (2014),
those of rice and wheat on Janni et al. (2006) and Di Giovanni et al. (2008),
and those of thale crest on Ferrari et al. (2003). Chr, chromosome.
Frontiers in Plant Science | www.frontiersin.org 3 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 1 | Treatments or stress stimuli affecting pgip expression in some plant species with a well characterized pgip family.
Pgip family Treatments or stress stimuli References
Rice Abscisic acid (ABA), brassinosteroid, gibberellic acid (GA), 3-indole acetic acid (IAA), jasmonic
acid (JA), kinetin, naphthalene acetic acid (NAA), salicylic acid (SA); Rhizoctonia solani
(necrotrophic fungus)
Janni et al., 2006; Lu et al., 2012
Wheat Bipolaris sorokiniana (necrotrophic fungus) and mechanical wounding Janni et al., 2013
Bean Oligogalacturonides (OGs); mechanical wounding; Botrytis cinerea, Sclerotinia sclerotiorum
(necrotrophic fungi); Colletotrichum lindemuthianum (hemibiotrophic fungus)
Bergmann et al., 1994; Nuss et al., 1996; Devoto
et al., 1997; D’Ovidio et al., 2004b; Oliveira et al.,
2010; Kalunke et al., 2011
Soybean Mechanical wounding; S. sclerotiorum (necrotrophic fungus) D’Ovidio et al., 2006; Kalunke et al., 2014
M. truncatula JA, SA, ABA; Colletotrichum trifolii (hemibiotrophic fungus) Song and Nam, 2005
Rapeseed JA, SA, mechanical wounding; S. sclerotiorum Hegedus et al., 2008
Pepper SA, Methyl jasmonate (Me-JA), ABA, wounding, cold treatment Wang et al., 2013
Arabidopsis OGs; JA; B. cinerea; Stemphylium solani (necrotrophic fungus); aluminum, low-pH, cold;
geminivirus
Ferrari et al., 2003; Ascencio-Ibanez et al., 2008;
Sawaki et al., 2009; Di et al., 2012; Kobayashi et al.,
2014
nature and are likely to have different specificities against micro-bial PGs, considering that single amino acid changes are able tochange specificity of the inhibitors (Leckie et al., 1999). Search-ing for PGIPs with novel specificities may allow to count ona much larger reservoir of possible genes for crop protection.A direct and simple strategy to isolate PGIPs with recognitioncapability against a given PG may be based on affinity chromath-ography methods, similar to that originally used to purify PGIPfrom P. vulgaris (Cervone et al., 1987), and mass spectrometry.Attempts to drive in vitro evolution of PGIPs to generate proteinswith improved inhibition properties have not been successful yet(Benedetti et al., 2011a).
The occurrence of PG-inhibiting activity in crude leaf proteinextracts of tetraploid wild wheat (T. dicoccoides) possessing nonfunctional pgip genes (Di Giovanni et al., 2008) suggested theexistance of pgip genes with a sequence divergent from the clas-sical one. This possibility, which deserves further investigation,is also supported by the finding that the wheat tissue containsPG-inhibiting proteins with N-terminal sequences (Lin and Li,2002; Kemp et al., 2003) different from TaPGIP1 and TaPGIP2(Janni et al., 2013) and from the pgip sequences reported sofar (http://www.ncbi.nlm.nih.gov/nucleotide/). Recently, a wheatgene with some sequence similarity to pgip genes has beenreported and was shown to be involved in the defense responseagainst Fusarium graminearum (Hou et al., 2014).
Structural Studies on the PG-PGIPInteraction
Thus, the possibility of engineering new forms of PGIPs dependson the detailed structural knowledge of the PG-PGIP interac-tion. Several structural studies have been performed (Mattei et al.,2001; King et al., 2002; Benedetti et al., 2011b, 2013; Gutierrez-Sanchez et al., 2012), but a high resolution 3D-structure of thePG-PGIP complex is still missing. The enzyme-inhibitor combi-nations that have been more extensively investigated, are thosethat PGIP2 from Phaseolus vulgaris (PvPGIP2) forms with PGfrom A. niger (AnPGII), F. phyllophilum (FpPG) and C. lupini(ClPG). Site-directed mutagenesis has shown that the residues
involved in the interaction are located in the concave surface ofthe inhibitor (Leckie et al., 1999; Federici et al., 2001; Spinelliet al., 2009; Benedetti et al., 2011b, 2013). Computational meth-ods such as the Codon Substitution Model in combination withthe Desolvation Energy Calculation and the Repeat Conserva-tion Mapping (RCM; Helft et al., 2011) have pinpointed severalresidues of PvPGIP2 responsible for the PG-inhibiting activity(Casasoli et al., 2009).
On the other hand, residues of PG that are critical for theinteraction with PGIP have been also studied. FvPG is 92.5%identical to FpPG, but is inhibited by neither PvPGIP2 nor otherknown PGIPs. By both loss- and gain-of-function site-directedmutations, a single amino acid at position 274 of both FvPGand FpPG was demonstrated to act as a switch for recognitionby PvPGIP2 (Raiola et al., 2008; Benedetti et al., 2013). Unfortu-nately, the lack of high-resolution structural information on thePG-PGIP complex does not allow to precisely identify the con-tacting residue in PGIP. Moreover, both PGs and PGIPs are gly-cosylated proteins (Caprari et al., 1993; Lim et al., 2009); however,whether glycosylation plays a role in the PGIP-PG interactionrequires further investigation. For example, glycosylation in pearlmillet PGIP was found to affect pH and temperature stability ofthe protein but not its capability of inhibiting AnPGII (Prabhuet al., 2015).
A single PGIP may display different mechanisms of PG inhi-bition (competitive, non competitive and mixed) suggesting thatthe protein is highly versatile in recognizing different epitopesof various PGs (Federici et al., 2001; King et al., 2002; Siciliaet al., 2005; Bonivento et al., 2008). Consequently, many 3D-models based on docking predictions have been proposed so far(Sicilia et al., 2005; Maulik et al., 2009; Prabhu et al., 2014). Tech-niques such as the mass amide exchange mass spectrometry inthe case of AnPGII and FpPG and the Small Angle X-ray Scat-tering (SAXS) in the case of FpPG and ClPG have producedmodels that, in some cases, are discordant. For example, while themass amide exchange mass spectrometry predicts that the area ofFpPG in contact with PvPGIP2 is located at the N-terminus andpredominantly on the underside of the enzyme beta-barrel struc-tures (Gutierrez-Sanchez et al., 2012), the SAXS analysis indicates
Frontiers in Plant Science | www.frontiersin.org 4 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 2 | Bulk PGIP purified from plants and tested against microbial PGs. These data update those reported in De Lorenzo et al. (2001).
Plant Tissue PGIP
preparation
Polygalacturonases References
Inhibited Not inhibited
Tomato (Solanum
lycopersici L.)
Stem Crude extract Ralstonia solanacearum Schacht et al., 2011
Tobacco (Nicotiana
tabacum L.)
Nectar Botrytis cinerea Thornburg et al., 2003
Potato (Solanum
tuberosum L.)
Gel
chromatography
Aspergillus niger Fusarium solani (isolate
3122)
Machinandiarena et al., 2001
Fusarium moniliforme§
Fusarilm solani isolate
1402
Common Bean
(Phaseolus vulgaris L.)
Leaves PG-Sepharose
chromatography
Fusarium anthophilum Fusarium verticillioides Raiola et al., 2008
Fusarium circinatum Fusarium proliferatum
ISPAVEmc 1189
Fusarium subglutinans. Fusarium nygamai
Fusarium proliferatum
isolate 1152
Fusarium proliferatum
PVS-Fu 64
Fusarium sacchari
Fusarium fujikuroi
F. thapsinum
Fusarium moniliforme§
FC-10
Fusarium moniliforme§
PD
Sella et al., 2004
Leek (Allium
ampeloprasum L.)
Basal leaves Mono-S
chromatography
Fusarium anthophilum Raiola et al., 2008
Fusarium circinatum
Fusarium subglutinans
Fusarium proliferatum
Fusarium sacchari
Fusarium fujikuroi
Fusarium verticillioides
Fusarium proliferatum
ISPAVEmc 1189
Fusarium nygamai
Asparagus (Asparagus
officinalis L.)
White spear Mono-S
chromatography
Fusarium anthophilum Raiola et al., 2008
Fusarium circinatum
Fusarium subglutinans.
Fusarium proliferatum
Fusarium sacchari
Fusarium fujikuroi
Fusarium verticillioides
Fusarium proliferatum
ISPAVEmc 1189
Fusarium nygamai
Pepper (Capsicum
annuum L.)
Fruit Ion-exchange
chromatography
Colletotrichum
gleosporoides,
Shivashankar et al., 2010
Colletotrichum capsici,
Colletotrichum
lindemuthianum
Sclerotium rolfsi
Fusarium moniliforme§
(Continued)
Frontiers in Plant Science | www.frontiersin.org 5 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 2 | Continued
Plant Tissue PGIP
preparation
Polygalacturonases References
Inhibited Not inhibited
Guava (Psidium
guajava L.)
Fruit Purified using a
Sephadex G-100
Aspergillus niger Deo and Shastri, 2003
“Oroblanco” grapefruit
hybrid (Citrus grandis × C.
paradisi Macf.)
Fruit Anion exchange
chromatography
Penicillium italicum D’hallewin et al., 2004
Botrytis cinerea
Apple (Malus
domestica L.)
Fruit Colletotrichum acutatum Gregori et al., 2008
Fruit skin Partial purified Botryosphaeria dothidea Glomerella cingulata Lee et al., 2006
Parenchymal
tissues
Partial purified Monilia fructigena Buza et al., 2004
Cantaloupe (Cucumis
melo L.)
Fruit Cation exchange
chromatography
Phomopsis cucurbitae Didymella bryoniae Fish and Davis, 2004
Aspergillus niger Rhizopus PG
Fusarium solani Fusarium verticillioides
Cotton (Gossypium
hirsutum L.)
Stem PG-affinity
chromatography
Aspergilus niger James and Dubery, 2001
Pear (Pyrus communis L.) Fruit Partial purified Verticillium dahliae Ladu et al., 2012; Faize
et al., 2003Botrytis cinerea
Venturia nashicola
Pearl millets (Pennisetum
glaucum (L) R. Br.)
Seedlings Crude extract Aspergilus niger Prabhu et al., 2012
Grass pea (Lathyrus
sativus L.)
Seeds Gel-filtration
chromatography
Aspergilus niger Tamburino et al., 2012
Rhizopus spp
Orange (Citrus reticulate
L.)
Fruit Partial purified Diaprepes abbreviatus Doostdar et al., 1997
Blue mustard (Chorispora
bungeana)
Leaves, stem,
root
Partial purified Aspergillus niger Di et al., 2009
Stemphylium solani
Ginseng
(Panax ginseng L.)
Crude extract Colletotrichum
gloeosporioides
Sathiyaraj et al., 2010
Phythium ultimum
Fusarium oxysporum
Rhizoctonia solani
Bread wheat (Triticum
aestivum L.)
Leaves Cation exchange
chromatography
Cochliobolus sativus Aspergillus niger (EPG I
and II)
Kemp et al., 2003
Cryphonectria
parasitica
Postia placenta
Fusarium moniliforme§
Colletotrichum
lindemuthianum
Aspergillus niger
exopolygalacturonase
Ralstonia
solanacearum
(Continued)
Frontiers in Plant Science | www.frontiersin.org 6 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 2 | Continued
Plant Tissue PGIP
preparation
Polygalacturonases References
Inhibited Not inhibited
Durum wheat
(Triticum turgidum ssp.
dicoccoides
Leaves Crude extract Fusarium graminearum Fusarium phyllophylum Janni et al., 2013
Bipolaris sorokiniana
Stenocarpella maydis
§Reclassified as Fusarium phyllophilum (Mariotti et al., 2008).
that the protein region in contact with PvPGIP2 is located atthe C-terminus of the enzyme and includes the loops surround-ing the active site cleft. A site-directed mutagenesis analysis hasbeen used to validate this second view (Benedetti et al., 2013).In general, low resolution techniques such as SAXS analysis ormass amide exchange mass spectrometry require validation bysite-directed mutagenesis to locate the contacting residues in aprotein complex.
The X-ray crystallography, successfully used to solve sev-eral high-resolution structures of PGs (van Santen et al., 1999;Federici et al., 2001; Bonivento et al., 2008) and that of PvPGIP2(Di Matteo et al., 2003), was so far unsuccessful in the caseof the PG-PGIP complex. This is probably due to the intrin-sic instability of the PG-PGIP interaction, which only occurs,under apoplastic conditions of pH and ionic strength, throughthe contact of only a few, sometimes only one, residues (Leckieet al., 1999). The use of a cross-linker for stabilizing the PG-PGIPcomplex coupled to techniques that allow the protein analysisdirectly in solution, such as SAXS and NMR spectroscopy (Wandand Englander, 1996; Nietlispach et al., 2004), may be a validalternative in order to obtain a detailed map of the contactingresidues but this requires a subsequent validation by site-directedmutagenesis.
PGIPs Engineered in Dicot Crops
The important role of PGIP in plant defense has been demon-strated by overexpressing pgip genes in several plant species. Inthese experiments, the source of the used genes was either thesame plant species utilized for transformation or a different one(Table 4). The transformation of the model plant A. thalianahas been particularly useful to highlight the potentiality of sev-eral pgip genes, namely the endogenous Atpgip1 and Atpgip2,the bean Pvpgip2 and the rapeseed (Brassica napus) Bnpgip1 orBnpgip2. Arabidopsis plants overexpressing Atpgip1 or Atpgip2showed a significant reduction of disease symptoms caused byB. cinerea (Ferrari et al., 2003) and were less susceptible againstthe hemibiotrophic fungal pathogen F. graminearum (Ferrariet al., 2012), the major causal agent of Fusarium head blight(FHB). Conversely, silencing of their expression using an anti-sense Atpgip, led to enhanced susceptibility (Ferrari et al., 2006).Arabidopsis plants expressing Pvpgip2, encoding an efficientinhibitor of the B. cinerea PG (ten Have et al., 1998), showedreduction of disease symptoms caused by B. cinerea and thoseexpressing the rapeseed genes Bnpgip1 and Bnpgip2 delayed thesymptoms caused by S. sclerotiorum (Bashi et al., 2013).
The protective potential of pgip genes has also been demon-strated in transgenic crops. The first transgenic crop plantobtained by using a pgip gene and tested against pathogenicmicroorganisms were tomatos expressing PvPGIP1 from P. vul-garis. These plants, however, did not show any increased resis-tance against Fusarium oxysporum f. sp. lycopersici, B. cinerea,and Alternaria solani. The negative result was due to the inabilityof PvPGIP1 to inhibit the PGs secreted by these fungi, as shownby in vitro inhibition assays and led to discovery of other forms ofPGIPs and eventually to the existence of a complex PGIP familyin French bean (Desiderio et al., 1997). A few years later, trans-genic tomato plants expressing a pear (Pyrus communis L.) PGIP(PcPGIP) capable of inhibiting the PGs secreted by B. cinerea,showed a reduction of disease lesions caused by this fungus bothon ripening fruit (15% reduction) and leaves (about 25% reduc-tion). The initial establishment of infection was not affected inthe transgenic plants but the later colonization of the host tissuewas significantly reduced (Powell et al., 2000).
Tobacco has been the most used crop plant for testing theeffect of PGIP expression on resistance to pathogens. Constitutiveand high-level expression of Pvpgip2 (from P. vulgaris), Vvpgip1(from V. vinifera), Capgip1 [from pepper (Capsicum annum)]and Brpgip2 (from B. rapa) have been obtained in transgenictobacco. Plants expressing PvPGIP2 showed about 35% reductionof symptoms caused by B. cinerea (Manfredini et al., 2005) and,more recently, were shown to display reduced disease symptomsagainst Rhizoctonia solani and two oomycete pathogens, Phy-tophthora parasitica var. nicotianae and the blue mold-causingagent Peronospora hyoscyami f. sp. tabacina (Borras-Hidalgoet al., 2012). Notably, the experiments against P. hyoscyami f.sp.tabacina were performed in the field during seasonal condi-tions that favor the pathogen spreading. In agreement with whatobserved under controlled conditions, resistance of transgenicplants was comparable to that exhibited by Nicotiana species(N. rustica, N. debneyi and N. megalosiphon) that are highlyresistant to blue mold disease. These transgenic plants express-ing PvPGIP2 represented the first example of PGIP-expressingplants subjected to field trails. Recently, transgenic rice express-ing OsPGIP1 showed also improved resistance against Rhizocto-nia solani in field experiments (Wang et al., 2014b).
Transgenic tobacco plants expressing the grapevine pgip geneVvpgip1 (Joubert et al., 2006) also showed a reduced (from 47to 69%) disease susceptibility to B. cinerea infection. As forplants expressing PvPGIP2, the resistance phenotype correlatedwith the accumulation of VvPGIP1 as well as with its capabil-ity of inhibiting the activity of PG secreted by B. cinerea, namely
Frontiers in Plant Science | www.frontiersin.org 7 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 3 | Pgip genes individually expressed in plants or in heterologous systems and tested for inhibition activity against microbial PGs.
Species Gene Heterologous systems Origin of purified PG References
Inhibited Not inhibited
Common bean (Phaseolus
vulgaris L.)
PvPGIP1 Transgenic tomato Fusarium oxysporum Desiderio et al., 1997
Botrytis cinerea
Alternaria solani
Stenocarpella maydis Berger et al., 2000
Aspergillus niger
PvPGIP1
PvPGIP2
PvPGIP3
PvPGIP4
PVX/Nicotiana benthamiana Aspergillus niger Lygus rugulipennis D’Ovidio et al., 2006; Frati
et al., 2006Fusarium moniliforme§ Adelphocoris lineolatus
Stenocarpella maydis Orthops kalmi
Colletotrichum acutatum Closterotomus norwegicus
Botrytis cinerea
PvPGIP2 Transgenic wheat Bipolaris sorokiniana Claviceps purpurea Janni et al., 2008; Volpi
et al., 2013F. graminearum
Transgenic Brassica napus Rhizoctonia solani Akhgari et al., 2012
Transgenic sugarbeet Fusarium phyllophilum FC10 Mohammadzadeh et al.,
2012
PVX/Nicotiana benthamiana Fusarium phyllophilum FC-10 Fusarium phyllophilum 25305 Mariotti et al., 2008
Fusarium phyllophilum 10241 Fusarium verticillioides 62264
Fusarium phyllophilum 25219 Fusarium verticillioides PD
Fusarium phyllophilum 25218
Runner bean (Phaseolus
coccineus L.)
PcPGIP2 PVX/Nicotiana benthamiana Fusarium moniliforme§ Farina et al., 2009
Aspergillus niger
Colletotrichum lupini
Botrytis cinerea
Tepary bean (Phaseolus
acutifolius L.)
PaPGIP2 PVX/Nicotiana benthamiana Fusarium moniliforme§ Farina et al., 2009
Aspergillus niger
Colletotrichum lupini
Botrytis cinerea
Lima bean (Phaseolus
lunatus L.)
PlPGIP2 PVX/Nicotiana benthamiana Fusarium moniliforme§ Farina et al., 2009
Aspergillus niger
Colletotrichum lupini
Botrytis cinerea
Soybean (Glycine max L.) GmPGIP1
GmPGIP2
PVX/Nicotiana benthamiana Sclerotinia sclerotiorum PGb D’Ovidio et al., 2006; Frati
et al., 2006Sclerotinia sclerotiorum PGa
Fusarium moniliforme§
Botrytis aclada
Aspergillus niger
Botrytis cinerea
Colletotrichum acutatum
Fusarium graminearum
Lygus rugulipennis
Adelphocoris lineolatus
Orthops kalmi
Closterotomus norwegicus
(Continued)
Frontiers in Plant Science | www.frontiersin.org 8 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 3 | Continued
Species Gene Heterologous systems Origin of purified PG References
Inhibited Not inhibited
GmPGIP3 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum PGb D’Ovidio et al., 2006; Frati
et al., 2006Sclerotinia sclerotiorum PGa
Fusarium moniliforme§
Botrytis aclada
Aspergillus niger
Botrytis cinerea
Colletotrichum acutatum
Fusarium graminearum
GmPGIP4 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum PGb D’Ovidio et al., 2006; Frati
et al., 2006Sclerotinia sclerotiorum PGa
Fusarium moniliforme§
Botrytis aclada
Aspergillus niger
Botrytis cinerea
Colletotrichum acutatum
Fusarium graminearum
GmPGIP7 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum
Fusarium graminearum Kalunke et al., 2014
Colletotrichum
acutatum
Aspergillus niger
Pepper (Capsicum
annum L.)
CaPGIP1
CaPGIP2
Escherichia coli Alternaria alternata Wang et al., 2013
Colletotrichum nicotianae
Rapeseed (Brassica
napus L.)
BnPGIP1 Pichia pastoris Sclerotinia sclerotiorum PG6 Bashi et al., 2013
Chinese cabbage (Brassica
rapa L.)
BrPGIP2 Transgenic Brassica rapa Pectobacterium carotovorum Hwang et al., 2010
Botryosphaeria dothidea
BrPGIP2 Escherichia coli Sclerotinia sclerotiorum HuangFu et al., 2014
Grapevine (Vitis vinifera L.) VvPGIP1 Transgenic tobacco Botrytis cinerea PGI Botrytis cinerea PG3 Joubert et al., 2006
Botrytis cinerea PG4 Aspergillus niger PGII
Botrytis cinerea PG6
Aspergillus. niger PGA
Aspergillus niger PGB
Aspergillus niger PGI Botrytis cinerea PG2 Joubert et al., 2007
Apple
(Malus domestica Borkh.)
MdPGIP1 Transgenic tobacco Colletotrichum lupini Aspergillus niger Oelofse et al., 2006
Botryosphaeria obtusa
Diaporthe ambigua
Transgenic potato Verticillium dahliae Gazendam et al., 2004
Pear (Pyrus communis L.) PpPGIP Transgenic grape Botrytis cinerea Agüero et al., 2005
Transgenic tomato Botrytis cinerea Powell et al., 2000
Transgenic persimmon Botrytis cinerea Tamura et al., 2004
(Continued)
Frontiers in Plant Science | www.frontiersin.org 9 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 3 | Continued
Species Gene Heterologous systems Origin of purified PG References
Inhibited Not inhibited
Raspberry (Rubus idaeus L.) RiPGIP Transgenic pea Stenocarpella maydis Richter et al., 2006
Colletotrichum lupini
Wheat
(Triticum aestivum L.)
TaPGIP1
TaPGIP2
PVX/Nicotiana benthamiana Fusarium phyllophylu Janni et al., 2013
Stenocarpella maydis
Bipolaris sorokiniana
Fusarium graminearum
Rice (Oryza sativa L.) OsPGIP1 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum Janni et al., 2006
Fusarium moniliforme§
Fusarium graminearum
Aspergillus niger
Botrytis cinerea
OsFOR1 Escherichia coli BL21 Aspergillus niger PG Jang et al., 2003
Pearl millet [Pennisetum
glaucum (L.) R. Br.]
PglPGIP1 Escherichia coli SHuffle® T7
Express
Aspergillus niger, AnPGII Fusarium moniliforme,
FmPGIII
Prabhu et al., 2014
Arabidopsis thaliana AtPGIP1
AtPGIP2
Transgenic Arabidopsis Colletotrichum gloeosporioides Aspergillus niger Frati et al., 2006; Ferrari
et al., 2012, 2003Stenocarpella maydis Fusarium moniliforme§
Botrytis cinerea Lygus rugulipennis
Fusarium graminearum Adelphocoris lineolatus
Orthops kalmi
Closterotomus norwegicus
§Reclassified as Fusarium phyllophilum FC10 (Mariotti et al., 2008).
BcPG1, BcPG3, and BcPG6. Several observations, however, sug-gest that PGIPmay improve resistance bymechanisms other thanclassical PGIP-PG inhibition. For example, non-infected trans-genic tobacco plants expressing Vvpgip1 show modified expres-sion patterns of genes involved in various metabolic pathways(Alexandersson et al., 2011) and an altered cell wall structure(Nguema-Ona et al., 2013). In these plants, lignin accumulationand arabinoxyloglucan-cellulose re-organization leads to a gen-eral strengthening/reinforcing of the cell wall that may contributeto an improved resistance against B. cinerea.
A reduction of disease symptoms (about 50%) caused byAlternaria alternata and Colletotrichum nicotianae was alsoobserved in transgenic tobacco lines expressing the pepperCaPGIP1 and, once again, resistance correlated with the inhi-bition capacity of purified CaPGIP1 against PG activity of bothfungal pathogens (Wang et al., 2013).
Within the Solanaceae family, transgenic potato (Solanumtuberosum) plants expressing the gene StPGIP1 from S. torvumshowed a 50% reduction of wilt disease symptoms caused byVerticillium dahliae and a normal plant growth (Guo et al.,2014). Transgenic potato plants overexpressing the apple pgip1gene showed protection against the same fungal pathogen butdisplayed an extended juvenile phase (Gazendam et al., 2004).
Transgenic grapevine (V. vinifera) plants constitutivelyexpressing the pear PcPGIP gene represent an interesting exam-ple of the potential of PGIP for protection against pathogens
other than fungi and oomycetes. These plants show a delayeddevelopment of the Pierce’s disease (PD) caused by bacterialpathogen Xylella fastidiosa (Agüero et al., 2005). Not only leafscorching and Xylella titre were reduced but also plants showeda better re-growth after pruning compared to infected untrans-formed controls. Moreover, an inverse dose-effect relationshipwas shown between development of PD and levels of PcPGIPactivity in the tissues. The improved resistance of the grapevineplants expressing PcPGIP against a bacterial pathogen was unex-pected, because until then the PGIP inhibition activity wasthought to be limited to fungal and insect PGs (Cervone et al.,1990; Johnston et al., 1993; D’Ovidio et al., 2004b). It was latershown that pear PcPGIP inhibits the PG encoded by X. fastid-iosa and that PG activity is a virulence factor of this pathogen(Roper et al., 2007; Pérez-Donoso et al., 2010). The observa-tion that PcPGIP is present in xylem exudates of non-transgenicscions grafted on transgenic rootstocks expressing PcPGIP sug-gests that grafting of non transgenic varieties on transgenic root-stocks represents, in this case, a useful agronomical practice forplant protection (Agüero et al., 2005).
The results obtained with X. fastidiosa prompted furtherinvestigations on the capability of PGIP of controlling bacte-rial diseases (summarized in Table 4). Transgenic tobacco plantsexpressing B. rapa BrPGIP2 were resistant against Pectobac-terium carotovorum, the causal agent of the soft rot disease,with a strong reduction (66–88%) of the symptoms as compared
Frontiers in Plant Science | www.frontiersin.org 10 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
TABLE 4 | List of transgenic crops produced using the gene coding for PGIP and their response to fungal, oomycetes or bacterial phytopathogens.
Transgenic crops PGIP genec Tested against fungal, oomycetes or bacterial
phytopathogens
References
Tomatoa (Solanum
lycopersicum L.)
PcPGIP Botrytis cinerea* Powell et al., 2000, 1994
PvPGIP1 Fusarium oxysporum f.sp. lycopersici† Desiderio et al., 1997
Botrytis cinerea†
Alternaria solani†
Tobaccoa (Nicotiana
tabacum L.)
PvPGIP2 Botrytis cinerea* Manfredini et al., 2005
Rhizoctonia solani* Borras-Hidalgo et al., 2012
Phytophthora parasitica*
Peronospora hyoscyami*
CaPGIP1 Alternaria alternata* Wang et al., 2013
Colletotrichum nicotianae*
VvPGIP1 Botrytis cinerea* Joubert et al., 2006
BrPGIP2 Pectobacterium carotovorum* Hwang et al., 2010
Potatoa (Solanum
tuberosum L.)
MdPGIP1 StPGIP Verticillium dahliae†
Verticillium dahliae*
Gazendam et al., 2004; Guo
et al., 2014
Brassica rapaa BrPGIP2 Pectobacterium carotovorum* Hwang et al., 2010
Rapeseeda
(Brassica napus L.)
BnPGIP2 Sclerotinia sclerotiorum* HuangFu et al., 2014
Peaa
(Pisum sativum L.)
RiPGIP Glomus intraradices9 Hassan et al., 2012
Grapevinea
(Vitis vinifera L.)
Ricea (Oriza sativa L.)
PcPGIP OsPGIP1 Botrytis cinerea* Agüero et al., 2005; Wang et al.,
2014bXylella fastidiosa*
Rhizoctonia solani
Wheatb
(Triticum aestivum L.,
Triticum durum Desf.)
PvPGIP2GmPGIP3 Bipolaris sorokiniana* Janni et al., 2008
Fusarium graminearum* Ferrari et al., 2012
Claviceps purpurea†
Bipolaris sorokiniana*
Gaeumannomyces graminis var. tritici*
Volpi et al., 2013; Wang et al.,
2014a
Arabidopsis thaliana
L.aPvPGIP2 Botrytis cinerea* Manfredini et al., 2005
AtPGIP1 AtPGIP2 Fusarium graminearum* Ferrari et al., 2012
BnPGIP1BnPGIP2 Sclerotinia sclerotiorum* Bashi et al., 2013
aThe transgenic gene was under control of CaMV 35S promoter.bThe transgenic gene was under control of Ubiquitin promoter.cPc, Pyrus communis; Pv, Phaseolus vulgaris; Ca, Capsicum annum; Vv, Vitis vinifera; Br, Brassica rapa; Md, Malus domestica; St, Solanum torvum; Ri, Rubus idaeus; Ac, Actinidia
deliciosa; At, Arabidopsis thaliana; Bn, Brassica napa.
*Showed enhanced resistance.†No evidence of enhanced resistance.9No effect on mycorrhization.
to wild-type plants (Hwang et al., 2010). The resistance cor-related with the inhibitory activity against P. carotovorum PGactivity found in the total protein extracts of the transgenicplants (Hwang et al., 2010). Also chinese cabbage (B. rapa ssp.pekinensis) plants overexpressing BrPGIP2 showed higher resis-tance against P. carotovorum and produced normal looking pods-like structures with no viable seeds. Combination of crossingwith non-transgenic plants did not restore fertility of the trans-genic plants, suggesting that mechanisms such as ploidy changesoccurring during the tissue culture stage or changes in cell-wallarchitecture of sexual organs are responsible for the abnormality(Hwang et al., 2010).
No phenotypic abnormalities were, instead, found in trans-genic tobacco plants expressing BrPGIP2 (Hwang et al., 2010),nor in rapeseed plants overexpressing the B. napus Bnpgip2. Thelatter plants displayed a significant reduction of rot caused by thenecrotrophic fungal pathogen S. sclerotiorum (HuangFu et al.,2014).
Additional PGIP-transgenic crops include pea (Pisumsativum L.), transformed with Ripgip from raspberry (Rubusidaeus L.) (Richter et al., 2006), persimmon (Diospyros kakiL.) and apple (Malus domestica Borkh.) transformed withpear PcPGIP (Szankowski et al., 2003; Tamura et al., 2004),sugarbeet (Beta vulgaris L.) transformed with bean Pvpgip2
Frontiers in Plant Science | www.frontiersin.org 11 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
(Mohammadzadeh et al., 2012), chickpea transformed witheither Ripgip or a pgip gene from kiwi fruit (Senthil et al., 2004),tobacco transformed with PpPGIP gene from Pyrus pyrifoliaNakai (Liu et al., 2013) and maize (Zea mays L.) transformedwith bean Pvpgip1 (O’Kennedy et al., 2001). The response ofthese plants to pathogens has not been reported yet. Transgenicpea plants expressing RiPGIP were instead evaluated for theirresponse to beneficial microorganisms. Glomus intraradices, anarbuscular mycorrhizal fungus, colonized roots of transgenicplants at an extend comparable to that observed in control nontransgenic plants, indicating that the expression of RiPGIP doesnot affect mycorrhization (Hassan et al., 2012).
PGIPs Engineered in Monocot Crops
Although the low pectin content of cereal species like wheatand rice indicates that this cell wall component may have amarginal role during infection, results show that the expres-sion of PGIP in transgenic plants limits some diseases caused byfungal pathogens (Janni et al., 2008; Ferrari et al., 2012; Wanget al., 2014a,b). In our labs, the bean Pvpgip2 gene was usedunder the constitutive promoter of the maize unbiquitin gene(Ubi-1) to transform both durum and bread wheat by particlebombardment. PvPGIP2 was correctly targeted to the apoplastand the transgenic plants did not show any major morpholog-ical and growth defects. Transgenic wheat showed a significantreduction (46–50%) of foliar spot blotch symptoms caused bythe hemibiotrophic fungal pathogen Bipolaris sorokiniana andimproved resistance (25–30%) against the hemibiotrophic fun-gal pathogen F. graminearum (Ferrari et al., 2012), the majorcausal agent of FHB in wheat. A reduced degradability of thetransgenic tissue by PG treatments correlated with the capacity ofPvPGIP2 to inhibit PG activity of B. sorokiniana and less stronglyPG of F. graminearum (Janni et al., 2008; Ferrari et al., 2012). Aninteresting aspect of the wheat plants expressing PvPGIP2 is that,under moderate infection with F. graminearum, the reduced FHBsymptoms are concomitant with a greater amount of total starchin the grains as compared to control plants (D’Ovidio et al., 2012).On the other hand, wheat plants expressing PvPGIP2 were sus-ceptible to the biotrophic fungal pathogen Claviceps purpurea,the causal agent of ergot disease probably because PvPGIP2 isnot able to inhibit the activity of C. purpurea CpPG1 and CpPG2(Volpi et al., 2013). Recently, transgenic wheat expressing thesoybean GmPGIP3 was shown to be resistant to both take-all andcommon root rot diseases caused by the fungal pathogen Gaeu-mannomyces graminis var. tritici and B. sorokiniana, respectively;symptoms were reduced of about 47–83% and 42–60%, respec-tively (Wang et al., 2014a). Similarly, the expression of OsPGIP1in transgenic rice enhanced resistence against Rhizoctonia solaniin field tests and resistance was related with the expression levelsof OsPGIP1 (Wang et al., 2014b).
Concluding Remarks and FutureChallenges
The results reported in this review clearly indicate that PGIP isuseful to improve resistance in different crop species. High-level
expression of PGIP does not prevent infection but limits sig-nificantly the colonization of the host tissue with a consequentpositive impact on crop yield and product quality. The efficacy ofPGIP to control diseases has been demonstrated against fungi,oomycetes and bacteria and is equally efficient against necro-rophic and hemibiotrophic pathogens. The experiments per-formed with biotrophs do not allow to draw any clear conclu-sion since the only fungal biotrophic pathogen analyzed, C. pur-pures, produced PG activity that was not inhibited by the PGIPexpressed in the transgenic plants (Volpi et al., 2013). The iden-tification and development of PGIPs with stronger and broaderinhibitory capacities may be useful to utilize these proteins incrop protection. Germplasm analysis to identify novel PGIPsis still limited (Farina et al., 2009) and the initial attemptsto drive in vitro evolution of PGIP to generate proteins withimproved inhibition properties have not been particularly suc-cessful (Benedetti et al., 2011a). Structural studies should beimplemented in order to obtain a detailed map of the contactsbetween various PGs and PGIPs. This is necessary not only forconstructing novel inhibitors with stronger activities but also forfuture programs of genome editing in which the existing genes ofa plant species may be ameliorated to better adapt to new virulentstrains of microorganisms evolving in nature.
The available results support the notion that inhibition ofthe microbial PG by PGIP is a prerequisite of the inhibitorsto confer resistance to transgenic plants against microbes. Thedelay of symptoms is often related to the capacity of PGIP toinhibit the PG activity secreted by the pathogens and, conse-quently, to reduce both tissue maceration and favor the releaseof OGs, as summarized in Figure 2. However, this aspect of thePGIP’s biology needs further investigation. In some cases PGIPhas been reported to confer resistance without any evidence ofPG-inhibition in vitro (Joubert et al., 2006). Moreover, some evi-dence suggests that the capability of reducing tissue maceration isassociated with the property of PGIP to bind pectin, likely shield-ing this component of the cell wall from PG activity (Spadoniet al., 2006). In this regard the observation that transgenic plantsexpressing PGIPs exhibit an altered gene expression and cell wallcomposition is also intriguing. It is not yet clear the mechanismthat links the ectopic expression of PGIP to alteration of geneexpression and whether this contributes to disease resistance(Alexandersson et al., 2011; Nguema-Ona et al., 2013).
An important but very little explored aspect of the PGIP biol-ogy is its possible role in processes of growth and development.Although plants overexpressing PGIPs do not show obviousmor-phological alterations, indeed several reports point to PGIP asa player in development. PGIP are induced, not only by phos-phate deficiency, but also by auxin treatment and in mutantsdefective in SIZ1, a SUMO (small ubiquitin-related modifier) E3ligase that is involved in several stress responses, including Pistarvation, and flowering (Sato and Miura, 2011). Suppression ofPGIPs under the control ABA insensitive 5 (ABI5) transcriptionfactor accompanies promotion of seed germination by the per-oxisomal ABC transporter PED3 (Kanai et al., 2010). Upregu-lation of PGIP2 correlates with the acquisition of competenceto form green callus in an auxin-rich callus induction medium(Che et al., 2007) and occurs in Arabidopsis tissue culture lines
Frontiers in Plant Science | www.frontiersin.org 12 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
FIGURE 2 | A model for the role of PGIP in the defense
response against pathogens. Delay of symptoms is related to the
inhibitory activity of PGIP toward PGs secreted by the pathogens
and likely to the accumulation of oligogalacturonide (OG) elicitors,
which are recognized by WAK1 and likely other receptors not yet
characterized. Cell wall modification and pectin shielding could also
play a role. Signaling cascades activated by OGs are described in
Ferrari et al. (2013).
in which the expression of the peroxidases PRX33 and PRX34is knocked down by antisense expression (O’Brien et al., 2012),whereas PGIP1 was identified in a proteomic study performed onArabidopsis etiolated hypocotyls used as a model of cells under-going elongation followed by growth arrest within a short time(Irshad et al., 2008). Finally, both PGIP1 and PGIP2 are associ-ated with cell wall stabilization at low pH under the control ofthe zinc-finger protein STOP1 (Sensitive to Proton Rhizotoxic-ity 1) and STOP2 (Kobayashi et al., 2014). A role of PGIP notonly in defense but also in growth and development implies that
the inhibitor may affect one or more of the many endogenousPGs expressed by plants. This is also an unexplored aspect of thePGIP biology and, at the moment, only one very old evidence isavailable showing that PGIPmay have a plant-derived PG partner(Cervone et al., 1990).
Acknowledgments
This research was supported by the Italian Ministry of Universityand Research (PRIN 2010–2011) to RD.
References
Agüero, C. B., Uratsu, S. L., Greve, C., Powell, A. L. T., Labavitch, J. M., Meredith,C. P., et al. (2005). Evaluation of tolerance to Pierce’s disease and Botrytis intransgenic plants of Vitis vinifera L. expressing the pear PGIP gene. Mol. Plant
Pathol. 6, 43–51. doi: 10.1111/j.1364-3703.2004.00262.xAkhgari, A., Motallebi, B., and Zamani, M. (2012). Bean polygalacturonase-
inhibiting protein expressed in transgenic Brassica napus inhibits poly-galacturonase from its fungal pathogen Rhizoctonia solani. Plant Prot. Sci.
48, 1–9.
Albersheim, P., and Anderson, A. J. (1971). Proteins from plant cell walls inhibitpolygalacturonases secreted by plant pathogens. Proc. Natl. Acad. Sci. U.S.A. 68,1815–1819.
Alexandersson, E., Becker, J. V. W., Jacobson, D., Nguema-Ona, E., Steyn,C., Denby, K. J., et al. (2011). Constitutive expression of a grapevinepolygalacturonase-inhibiting protein affects gene expression and cell wall prop-erties in uninfected tobacco. BMC Res. Notes 4:493. doi: 10.1186/1756-0500-4-493
Ascencio-Ibanez, J. T., Sozzani, R., Lee, T.-J., Chu, T.-M., Wolfinger, R. D., Cella,R., et al. (2008). Global analysis of Arabidopsis gene expression uncovers a
Frontiers in Plant Science | www.frontiersin.org 13 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
complex array of changes impacting pathogen response and cell cycle duringgeminivirus infection. Plant Physiol. 148, 436–454. doi: 10.1104/pp.108.121038
Bashi, Z. D., Rimmer, S. R., Khachatourians, G. G., and Hegedus, D. D. (2013).Brassica napus polygalacturonase inhibitor proteins inhibit Sclerotinia sclero-
tiorum polygalacturonase enzymatic and necrotizing activities and delay symp-toms in transgenic plants. Can. J. Microbiol. 59, 79–86. doi: 10.1139/cjm-2012-0352
Benedetti, M., Andreani, F., Leggio, C., Galantini, L., Di Matteo, A., Pavel, N. V.,et al. (2013). A single amino-acid substitution allows endo-polygalacturonaseof Fusarium verticillioides to acquire recognition by PGIP2 from Phaseolus
vulgaris. PLoS ONE 8:e80610. doi: 10.1371/journal.pone.0080610Benedetti, M., Bastianelli, E., Salvi, G., De Lorenzo, G., and Caprari, C.
(2011a). Artificial evolution corrects a repulsive amino acid in polygalac-turonase inhibiting proteins (PGIPs). J. Plant Pathol. 93, 89–95. doi:10.4454/jpp.v93i1.277
Benedetti, M., Leggio, C., Federici, L., De Lorenzo, G., Pavel, N. V., and Cervone,F. (2011b). Structural resolution of the complex between a fungal polygalactur-onase and a plant polygalacturonase-inhibiting protein by small-angle X-rayscattering. Plant Physiol. 157, 599–607. doi: 10.1104/pp.111.181057
Berger, D. K., Oelofse, D., Arendse, M. S., Du Plessis, E., and Dubery, I. A. (2000).Bean polygalacturonase inhibitor protein-1 (PGIP-1) inhibits polygalactur-onases from Stenocarpella maydis. Physiol. Mol. Plant Pathol. 57, 5–14. doi:10.1006/pmpp.2000.0274
Bergmann, C. W., Ito, Y., Singer, D., Albersheim, P., Darvill, A. G., Benhamou,N., et al. (1994). Polygalacturonase-inhibiting protein accumulates in Phaseo-
lus vulgaris L. in response to wounding, elicitors and fungal infection. Plant J.5, 625–634. doi: 10.1111/j.1365-313X.1994.00625.x
Bonivento, D., Pontiggia, D., Di Matteo, A., Fernandez-Recio, J., Salvi, G.,Tsernoglou, D., et al. (2008). Crystal structure of the endopolygalactur-onase from the phytopathogenic fungus Colletotrichum lupini and its inter-action with polygalacturonase inhibiting proteins. Proteins 70, 294–299. doi:10.1002/prot.21610
Borras-Hidalgo, O., Caprari, C., Hernandez-Estevez, I., De Lorenzo, G., and Cer-vone, F. (2012). A gene for plant protection: expression of a bean polygalac-turonase inhibitor in tobacco confers a strong resistance against Rhizocto-
nia solani and two oomycetes. Front. Plant Sci. 3:268. doi: 10.3389/fpls.2012.00268
Buza, N. L., Krinitsyna, A. A., Protsenko,M. A., andVartapetyan, V. V. (2004). Roleof the polygalacturonidase inhibitor Protein in the ripening of apples and theirresistance to Monilia fructigena, a causative agent of fruit rot. Appl. Biochem.
Microbiol. 40, 89–92. doi: 10.1023/B:ABIM.0000010361.48129.6eCantu, D., Vicente, A. R., Labavitch, J. M., Bennett, A. B., and Powell, A. L. (2008).
Strangers in the matrix: plant cell walls and pathogen susceptibility. TrendsPlant Sci. 13, 610–617. doi: 10.1016/j.tplants.2008.09.002
Caprari, C., Bergmann, C., Migheli, Q., Salvi, G., Albersheim, P., Darvill, A., et al.(1993). Fusarium moniliforme secretes four endopolygalacturonases derivedfrom a single gene product. Physiol. Mol. Plant Pathol. 43, 453–462. doi:10.1006/pmpp.1993.1073
Casasoli, M., Federici, L., Spinelli, F., Di Matteo, A., Vella, N., Scaloni, F., et al.(2009). Integration of evolutionary and desolvation energy analysis identifiesfunctional sites in a plant immunity protein. Proc. Natl. Acad. Sci. U.S.A. 106,7666–7671. doi: 10.1073/pnas.0812625106
Cervone, F., De Lorenzo, G., Degrà, L., Salvi, G., and Bergami, M. (1987). Purifi-cation and characterization of a polygalacturonase-inhibiting protein fromPhaseolus vulgaris L. Plant Physiol. 85, 631–637. doi: 10.1104/pp.85.3.631
Cervone, F., De Lorenzo, G., Pressey, R., Darvill, A. G., and Albersheim, P.(1990). Can Phaseolus PGIP inhibit pectic enzymes from microbes and plants?Phytochemistry 29, 447–449. doi: 10.1016/0031-9422(90)85094-V
Cervone, F., Hahn, M. G., De Lorenzo, G., Darvill, A., and Albersheim, P. (1989).Host-pathogen interactions. XXXIII. A plant protein converts a fungal patho-genesis factor into an elicitor of plant defense responses. Plant Physiol. 90,542–548. doi: 10.1104/pp.90.2.542
Che, P., Lall, S., and Howell, S. H. (2007). Developmental steps in acquiringcompetence for shoot development in Arabidopsis tissue culture. Planta 226,1183–1194. doi: 10.1007/s00425-007-0565-4
Chen, X., Liu, X., Zuo, S., Ma, Y., Tong, Y., Pan, X., et al. (2011). Prokaryoticexpression of riceOspgip1 gene and bioinformatic analysis of encoded product.Rice Sci. 18, 250–256. doi: 10.1016/S1672-6308(12)60002-X
D’hallewin, G., Schirra, M., Powell, A. L., Greve, L. C., and Labavitch, J. M. (2004).Properties of a polygalacturonase-inhibiting protein isolated from ’Oroblanco’grapefruit. Physiol. Plant. 120, 395–404. doi: 10.1111/j.0031-9317.2004.00264.x
D’Ovidio, R., Laino, P., Janni, M., Botticella, E., Di Carli, M. S., Benvenuto,E., et al. (2012). “Proteomic analysis of mature kernels of Fusarium gramin-
earum-infected transgenic bread wheat expressing PGIP,” in Proceedings of 11thInternational Gluten Workshop, eds Z. He and D. Wang (Beijing), 18.
D’Ovidio, R., Mattei, B., Roberti, S., and Bellincampi, D. (2004a). Polygalac-turonases, polygalacturonase-inhibiting proteins and pectic oligomers inplant-pathogen interactions. Biochim. Biophys. Acta 1696, 237–244. doi:10.1016/j.bbapap.2003.08.012
D’Ovidio, R., Raiola, A., Capodicasa, C., Devoto, A., Pontiggia, D., Roberti,S., et al. (2004b). Characterization of the complex locus of bean encod-ing polygalacturonase-inhibiting proteins reveals subfunctionalization fordefense against fungi and insects. Plant Physiol. 135, 2424–2435. doi:10.1104/pp.104.044644
D’Ovidio, R., Roberti, S., Giovanni, M. D., Capodicasa, C., Melaragni, M., Sella, L.,et al. (2006). The characterization of the soybean polygalacturonase-inhibitingproteins (Pgip) gene family reveals that a single member is responsible for theactivity detected in soybean tissues. Planta 224, 633–645. doi: 10.1007/s00425-006-0235-y
De Lorenzo, G., D’Ovidio, R., and Cervone, F. (2001). The role ofpolygalacturonase-inhibiting proteins (PGIPs) in defense against pathogenicfungi. Annu. Rev. Phytopathol. 39, 313–335. doi: 10.1146/annurev.phyto.39.1.313
De Lorenzo, G., and Ferrari, S. (2002). Polygalacturonase-inhibiting proteins indefense against phytopathogenic fungi. Curr. Opin. Plant Biol. 5, 295–299. doi:10.1016/S1369-5266(02)00271-6
Deo, A., and Shastri, N. V. (2003). Purification and characterization ofpolygalacturonase-inhibitory proteins from Psidium guajava Linn. (guava)fruit. Plant Sci. 164, 147–156. doi: 10.1016/S0168-9452(02)00337-0
Desiderio, A., Aracri, B., Leckie, F., Mattei, B., Salvi, G., Tigelaar, H., et al. (1997).Polygalacturonase-inhibiting proteins (PGIPs) with different specificities areexpressed in Phaseolus vulgaris. Mol. Plant Microbe Interact. 10, 852–860. doi:10.1094/MPMI.1997.10.7.852
Devoto, A., Clark, A. J., Nuss, L., Cervone, F., and De Lorenzo, G.(1997). Developmental and pathogen-induced accumulation of transcriptsof polygalacturonase-inhibiting protein in Phaseolus vulgaris L. Planta 202,284–292. doi: 10.1007/s004250050130
Di, C., Li, M., Long, F., Bai, M., Zheng, X., Xu, S., et al. (2009). Molec-ular cloning, functional analysis and localization of a novel gene encod-ing polygalacturonase-inhibiting protein in Chorispora bungeana. Planta 231,169–178. doi: 10.1007/s00425-009-1039-7
Di, C. X., Zhang, H., Sun, Z. L., Jia, H. L., Yang, L. N., Si, J., et al. (2012). Spatialdistribution of polygalacturonase-inhibiting proteins in Arabidopsis and theirexpression induced by Stemphylium solani infection. Gene 506, 150–155. doi:10.1016/j.gene.2012.06.085
Di Giovanni, M., Cenci, A., Janni, M., and D’Ovidio, R. (2008). A LTR copia retro-transposon andMutator transposons interrupt pgip genes in cultivated andwildwheats. Theor. Appl. Genet. 116, 859–867. doi: 10.1007/s00122-008-0719-1
Di Matteo, A., Bonivento, D., Tsernoglou, D., Federici, L., and Cervone, F. (2006).Polygalacturonase-inhibiting protein (PGIP) in plant defence: a structural view.Phytochemistry 67, 528–533. doi: 10.1016/j.phytochem.2005.12.025
Di Matteo, A., Federici, L., Mattei, B., Salvi, G., Johnson, K. A., Savino, C., et al.(2003). The crystal structure of polygalacturonase-inhibiting protein (PGIP),a leucine-rich repeat protein involved in plant defense. Proc. Natl. Acad. Sci.U.S.A. 100, 10124–10128. doi: 10.1073/pnas.1733690100
Doostdar, H., McCollum, T. G., and Mayer, R. T. (1997). Purification and char-acterization of an endo-polygalacturonase from the gut of West Indies sugar-cane rootstalk borer weevil (Diaprepes abbreviatus L.) larvae. Comp. Biochem.
Physiol. Part B Biochem. Mol. Bio. 118, 861–867.Faize,M., Sugiyama, T., Faize, L., and Ishii, H. (2003). Polygalacturonase-inhibiting
protein (PGIP) from Japanese pear: possible involvement in resistance againstscab. Physiol. Mol. Plant Pathol. 63, 319–327. doi: 10.1016/j.pmpp.2004.03.006
Farina, A., Rocchi, V., Janni, M., Benedettelli, S., De Lorenzo, G., and D’Ovidio, R.(2009). The bean polygalacturonase-inhibiting protein 2 (PvPGIP2) is highlyconserved in common bean (Phaseolus vulgaris L.) germplasm and relatedspecies. Theor. Appl. Genet. 118, 1371–1379. doi: 10.1007/s00122-009-0987-4
Frontiers in Plant Science | www.frontiersin.org 14 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
Federici, L., Caprari, C., Mattei, B., Savino, C., Di Matteo, A., De Lorenzo, G., et al.(2001). Structural requirements of endopolygalacturonase for the interactionwith PGIP (polygalacturonase-inhibiting protein). Proc. Natl. Acad. Sci. U.S.A.98, 13425–13430. doi: 10.1073/pnas.231473698
Federici, L., Di Matteo, A., Fernandez-Recio, J., Tsernoglou, D., and Cervone, F.(2006). Polygalacturonase inhibiting proteins: players in plant innate immu-nity? Trends Plant Sci. 11, 65–70. doi: 10.1016/j.tplants.2005.12.005
Ferrari, S., Galletti, R., Vairo, D., Cervone, F., and De Lorenzo, G. (2006).Antisense expression of the Arabidopsis thaliana AtPGIP1 gene reducespolygalacturonase-inhibiting protein accumulation and enhances suscepti-bility to Botrytis cinerea. Mol. Plant Microbe Interact. 19, 931–936. doi:10.1094/MPMI-19-0931
Ferrari, S., Savatin, D. V., Sicilia, F., Gramegna, G., Cervone, F., and DeLorenzo, G. (2013). Oligogalacturonides: plant damage-associated molecularpatterns and regulators of growth and development. Front. Plant Sci. 4:49. doi:10.3389/fpls.2013.00049
Ferrari, S., Sella, L., Janni, M., De Lorenzo, G., Favaron, F., and D’Ovidio, R. (2012).Transgenic expression of polygalacturonase-inhibiting proteins in Arabidopsis
and wheat increases resistance to the flower pathogen Fusarium graminearum.Plant Biol. 14, 31–38. doi: 10.1111/j.1438-8677.2011.00449.x
Ferrari, S., Vairo, D., Ausubel, F. M., Cervone, F., and De Lorenzo, G.(2003). Tandemly duplicated Arabidopsis genes that encode polygalacturonase-inhibiting proteins are regulated coordinately by different signal transductionpathways in response to fungal infection. Plant Cell Online 15, 93–106. doi:10.1105/tpc.005165
Fish,W.W., andDavis, A. R. (2004). The purification, physical/chemical character-ization, and cDNA sequence of cantaloupe fruit polygalacturonase-inhibitingprotein. Phytopathology 94, 337–344. doi: 10.1094/PHYTO.2004.94.4.337
Frati, F., Galletti, R., De Lorenzo, G., Salarino, G., and Conti, E. (2006). Activity ofendo-polygalacturonases in mirid bugs (Heteroptera: Miridae) and their inhi-bition by plant cell wall proteins (PGIPs). Eur. J. Entomol. 103, 515–522. doi:10.14411/eje.2006.067
Gazendam, I., Oelofse, D., and Berger, D. K. (2004). High-level expression of applePGIP1 is not sufficient to protect transgenic potato againstVerticillium dahliae.Physiol. Mol. Plant Pathol. 65, 145–155. doi: 10.1016/j.pmpp.2005.01.002
Gomathi, V., and Gnanamanickam, S. S. (2004). Polygalacturonase-inhibitingproteins in plant defence. Curr. Sci. 87, 1211–1217.
Gregori, R., Mari, M., Bertolini, P., Barajas, J. A., Tian, J. B., and Labavitch, J. M.(2008). Reduction of Colletotrichum acutatum infection by a polygalacturonaseinhibitor protein extracted from apple. Postharvest. Biol. Technol. 48, 309–313.doi: 10.1016/j.postharvbio.2007.10.006
Guo, J.-L., Zhu, Y.-P., Shi, K., Jue, D.-W., Liu, S.-P., Hong, Y.-B., et al. (2014).Over-expression of Solanum torvum PGIP enhances resistance to Verticillium
dahliae in transgenic potato plants. Bothalia Pretoria. 44, 392–404.Gutierrez-Sanchez, G., King, D., Kemp, G., and Bergmann, C. (2012). SPR and
differential proteolysis/MS provide further insight into the interaction betweenPGIP2 and EPGs. Fungal Biol. 116, 737–746. doi: 10.1016/j.funbio.2012.04.010
Hassan, F., Noorian, M. S., and Jacobsen, H. J. (2012). Effect of antifungal genesexpressed in transgenic pea (Pisum sativum L.) on root colonization withGlomus intraradices. GM Crops Food 3, 301–309. doi: 10.4161/gmcr.21897
Hegedus, D. D., Li, R., Buchwaldt, L., Parkin, I., Whitwill, S., Coutu, C., et al.(2008). Brassica napus possesses an expanded set of polygalacturonase inhibitorprotein genes that are differentially regulated in response to Sclerotinia scle-
rotiorum infection, wounding and defense hormone treatment. Planta 228,241–253. doi: 10.1007/s00425-008-0733-1
Helft, L., Reddy, V., Chen, X., Koller, T., Federici, L., Recio, J. F., et al. (2011). LRRConservation mapping to predict functional sites within protein leucine-richrepeat domains. PLoS ONE 6:e21614. doi: 10.1371/journal.pone.0021614
Hou, W., Mu, J., Li, A., Wang, H., and Kong, L. (2014). Identification ofa wheat polygalacturonase-inhibiting protein involved in Fusarium headblight resistance. Eur. J. Plant Pathol. 141, 731–745. doi: 10.1007/s10658-014-0574-7
HuangFu, H., Guan, C., Jin, F., and Yin, C. (2014). Prokaryotic expression andprotein function of Brassica napus PGIP2 and its genetic transformation. PlantBiotechnol. Rep. 8, 171–181. doi: 10.1007/s11816-013-0307-y
Hwang, B. H., Bae, H., Lim, H. S., Kim, K. B., Kim, S. J., Im, et al. (2010). Overex-pression of polygalacturonase-inhibiting protein 2 (PGIP2) of Chinese cabbage(Brassica rapa ssp. pekinensis) increased resistance to the bacterial pathogen
Pectobacterium carotovorum ssp. carotovorum. Plant Cell Tiss. Organ Cult. 103,293–305. doi: 10.1007/s11240-010-9779-4
Irshad, M., Canut, H., Borderies, G., Pont-Lezica, R., and Jamet, E. (2008). Anew picture of cell wall protein dynamics in elongating cells of Arabidop-sis thaliana: Confirmed actors and newcomers. BMC Plant Biol. 8:94. doi:10.1186/1471-2229-8-94
James, J. T., and Dubery, I. A. (2001). Inhibition of polygalacturonase fromVerticillium dahliae by a polygalacturonase inhibiting protein from cotton.Phytochemistry 57, 149–156. doi: 10.1016/S0031-9422(01)00024-3
Jang, S., Lee, B., Kim, C., Kim, S. J., Yim, J., Han, J. J., et al. (2003). TheOsFOR1 gene encodes a polygalacturonase-inhibiting protein (PGIP) thatregulates floral organ number in rice. Plant Mol. Biol. 53, 357–372. doi:10.1023/B:PLAN.0000006940.89955.f1
Janni, M., Bozzini, T., Moscetti, I., Volpi, C., and D’Ovidio, R. (2013). Func-tional characterisation of wheat Pgip genes reveals their involvement in thelocal response to wounding. Plant Biol. Stuttg. Ger. 15, 1019–1024. doi:10.1111/plb.12002
Janni, M., Giovanni, M., Roberti, S., Capodicasa, C., and D’Ovidio, R. (2006). Char-acterization of expressed Pgip genes in rice and wheat reveals similar extentof sequence variation to dicot PGIPs and identifies an active PGIP lacking anentire LRR repeat. Theor. Appl. Genet. 113, 1233–1245. doi: 10.1007/s00122-006-0378-z
Janni, M., Sella, L., Favaron, F., Blechl, A. E., De Lorenzo, G., and D’Ovidio, R.(2008). The expression of a bean PGIP in transgenic wheat confers increasedresistance to the fungal pathogen Bipolaris sorokiniana. Mol. Plant Microbe
Interact. 21, 171–177. doi: 10.1094/MPMI-21-2-0171Johnston, D. J., Ramanathan, V., and Williamson, B. (1993). A protein from
immature raspberry fruits which inhibits endopolygalacturonases from Botry-
tis cinerea and other micro-organisms. J. Exp. Bot. 44, 971–976. doi:10.1093/jxb/44.5.971
Joubert, D. A., Kars, I., Wagemakers, L., Bergmann, C., Kemp, G., Vivier, M. A.,et al. (2007). A polygalacturonase-inhibiting protein from grapevine reducesthe symptoms of the endopolygalacturonase BcPG2 from Botrytis cinerea inNicotiana benthamiana leaves without any evidence for in vitro interaction.Mol. Plant Microbe Interact. 20, 392–402. doi: 10.1094/MPMI-20-4-0392
Joubert, D. A., Slaughter, A. R., Kemp, G., Becker, J. V. W., Krooshof, G. H.,Bergmann, C., et al. (2006). The grapevine polygalacturonase-inhibiting protein(VvPGIP1) reduces Botrytis cinerea susceptibility in transgenic tobacco and dif-ferentially inhibits fungal polygalacturonases. Transgenic Res. 15, 687–702. doi:10.1007/s11248-006-9019-1
Kalunke, R. M., Cenci, A., Volpi, C., O’Sullivan, D. M., Sella, L., Favaron, F.,et al. (2014). The pgip family in soybean and three other legume species: evi-dence for a birth-and-death model of evolution. BMC Plant Biol. 14:189. doi:10.1186/s12870-014-0189-3
Kalunke, R. M., Janni, M., Sella, L., David, P., Geffroy, V., Favaron, F., et al. (2011).Transcript analysis of the bean polygalacturonase inhibiting protein gene fam-ily reveals that Pvpgip2 is expressed in the whole plant and is strongly inducedby pathogen infection. J. Plant Pathol. 93, 141–148. doi: 10.4454/jpp.v93i1.284
Kanai, M., Nishimura, M., and Hayashi, M. (2010). A peroxisomal ABC trans-porter promotes seed germination by inducing pectin degradation under thecontrol of ABI5. Plant J. 62, 936–947. doi: 10.1111/j.1365-313X.2010.04205.x
Kemp, G., Bergmann, C. W., Clay, R., Van der Westhuizen, A. J., andPretorius, Z. A. (2003). Isolation of a polygalacturonase-inhibiting pro-tein (PGIP) from wheat. Mol. Plant Microbe Interact. 16, 955–961. doi:10.1094/MPMI.2003.16.11.955
King, D., Bergmann, C., Orlando, R., Benen, J. A., Kester, H. C., and Visser,J. (2002). Use of amide exchange mass spectrometry to study confor-mational changes within the endopolygalacturonase II-homogalacturonan-polygalacturonase inhibiting protein system. Biochemistry 41, 10225–10233.doi: 10.1021/bi020119f
Kirsch, R., Wielsch, N., Vogel, H., Svatoš A., Heckel, D. G., and Pauchet, Y.(2012). Combining proteomics and transcriptome sequencing to identify activeplant-cell-wall-degrading enzymes in a leaf beetle. BMC Genomics 13:587. doi:10.1186/1471-2164-13-587
Kobayashi, Y., Ohyama, Y., Kobayashi, Y., Ito, H., Iuchi, S., Fujita, M., et al.(2014). STOP2 activates transcription of several genes for Al- and low pH-tolerance that are regulated by STOP1 in Arabidopsis. Mol. Plant 7, 311–322.doi: 10.1093/mp/sst116
Frontiers in Plant Science | www.frontiersin.org 15 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
Ladu, G., Pani, G., Venditti, T., Dore, A., Molinu,M. G., andD’Hallewin, G. (2012).Natural resistance against pre- and post-harvest pathogens in Sardinian pearsgermoplasm. Commun. Agric. Appl. Biol. Sci. 77, 163–171.
Lagaert, S., Beliën, T., and Volckaert, G. (2009). "Plant cell walls: protecting thebarrier from degradation by microbial enzymes" Semin. Cell Dev. Biol. 20,1064–1073. doi: 10.1016/j.semcdb.2009.05.008
Leckie, F., Mattei, B., Capodicasa, C., Hemmings, A., Nuss, L., Aracri, B., et al.(1999). The specificity of polygalacturonaseinhibiting protein (PGIP): a singleamino acid substitution in the solvent-exposed b-strand/b-turn region of theleucine-rich repeats (LRRs) confers a new recognition capability. EMBO J. 18:2352–2363 doi: 10.1093/emboj/18.9.2352
Lee, H. D., Bae, H., Kang, I. K., Byun, J. K., and Kang, S. G. (2006). Characteri-zation of an apple polygalacturonase-inhibiting protein (PGIP) that specificallyinhibits an endopolygalacturonase (PG) purified from apple fruits infected withBotryosphaeria dothidea. J. Microbio. Biotech. 16, 1192–1200.
Lim, J. M., Aoki, K., Angel, P., Garrison, D., King, D., Tiemeyer, M., et al. (2009).Mapping glycans onto specific N-linked glycosylation sites of Pyrus commu-
nis PGIP redefines the interface for EPG-PGIP interactions. J. Proteome Res. 8,673–680. doi: 10.1021/pr800855f
Lin, W., and Li, Z. (2002). The partial structure of wheat polygalacturonaseinhibiting protein. Chin. J. Biochem. Mol. Bio. 18, 197–201.
Liu, D., Li, W., He, X., Ding, Y., Chen, C., and Ge, F. (2013). Characterization andfunctional analysis of a novel PGIP gene from Pyrus pyrifoliaNakai cv Huobali.Acta Physiol. Plant. 35, 1247–1256. doi: 10.1007/s11738-012-1164-y
Lu, L., Zhou, F., Zhou, Y., Fan, X., Ye, S., Wang, L., et al. (2012). Expressionprofile analysis of the polygalacturonase-inhibiting protein genes in rice andtheir responses to phytohormones and fungal infection. Plant Cell Rep. 31,1173–1187. doi: 10.1007/s00299-012-1239-7
Machinandiarena, M. F., Olivieri, F. P., Daleo, G. R., and Oliva, C. R. (2001).Isolation and characterization of a polygalacturonase-inhibiting protein frompotato leaves. Accumulation in response tosalicylic acid, wounding andinfection. Plant Physiol. Biochem. 39, 129–136. doi: 10.1016/S0981-9428(00)01228-6
Manfredini, C., Sicilia, F., Ferrari, S., Pontiggia, D., Salvi, G., Caprari, C., et al.(2005). Polygalacturonase-inhibiting protein 2 of Phaseolus vulgaris inhibitsBcPG1, a polygalacturonase of Botrytis cinerea important for pathogenicity,and protects transgenic plants from infection. Physiol. Mol. Plant Pathol. 67,108–115. doi: 10.1016/j.pmpp.2005.10.002
Mariotti, L., Casasoli, M., Migheli, Q., Balmas, V., Caprari, C., and DeLorenzo, G. (2008). Reclassification of Fusarium verticillioides (syn. F. monil-
iforme) strain FC-10 as F. phyllophilum. Mycol. Res. 112, 1010–1011. doi:10.1016/j.mycres.2008.07.004
Mattei, B., Bernalda, M. S., Federici, L., Roepstorff, P., Cervone, F., and Boffi,A. (2001). Secondary structure and post-translational modifications of theleucine-rich repeat protein PGIP (polygalacturonase-inhibiting protein) fromPhaseolus vulgaris. Biochemistry 40, 569–576. doi: 10.1021/bi0017632
Maulik, A., Ghosh, H., and Basu, S. (2009). Comparative study of protein-proteininteraction observed in Polygalacturonase-inhibiting proteins from Phaseolus
vulgaris and Glycine max and Polygalacturonase from Fusarium moniliforme.BMC Genomics 10:S19. doi: 10.1186/1471-2164-10-S3-S19
Michelmore, R. W., andMeyers, B. C. (1998). Clusters of resistance genes in plantsevolve by divergent selection and a birth-and-death process. Genome Res. 8,1113–1130.
Misas-Villamil, J. C., and van der Hoorn, R. A. (2008). Enzyme-inhibitor interac-tions at the plant-pathogen interface. Curr. Opin. Plant Biol. 11, 380–388. doi:10.1016/j.pbi.2008.04.007
Mohammadzadeh, R., Zamani, M., Motallebi, M., Norouzi, P., Jourabchi, E.,Benedetti, M., et al. (2012). Agrobacterium tumefaciens-mediated introduc-tion of polygalacturonase inhibiting protein 2 gene (PvPGIP2) from Phaseolusvulgaris into sugar beet (Beta vulgaris L.). Aust. J. Crop Sci. 6, 1290–1297.
Nguema-Ona, E., Moore, J. P., Fagerström, A. D., Fangel, J. U., Willats, W. G.,Hugo, A., et al. (2013). Overexpression of the grapevine PGIP1 in tobaccoresults in compositional changes in the leaf arabinoxyloglucan network inthe absence of fungal infection. BMC Plant Biol. 13:46. doi: 10.1186/1471-2229-13-46
Nietlispach, D., Mott, H. R., Stott, K. M., Nielsen, P. R., Thiru, A., and Laue, E. D.(2004). Structure determination of protein complexes by NMR. Methods Mol.
Biol. 278, 255–288. doi: 10.1385/1-59259-809-9:255
Nuss, L., Mahe, A., Clark, A. J., Grisvard, J., Dron, M., Cervone, F., et al.(1996). Differential accumulation of PGIP (polygalacturonase inhibiting pro-tein) mRNA in two near-isogenic lines of Phaseolus vulgaris L. upon infectionwith Colletotrichum lindemuthianum. Physiol. Mol. Plant Pathol. 48, 83–89.
O’Kennedy, M., Burger, J., and Berger, D. (2001). Transformation of elitewhite maize using the particle inflow gun and detailed analysis of a low-copy integration event. Plant Cell Rep. 20, 721–730. doi: 10.1007/s002990100383
O’Brien, J. A., Daudi, A., Finch, P., Butt, V. S., Whitelegge, J. P., Souda, P.,et al. (2012). A peroxidase-dependent apoplastic oxidative burst in culturedarabidopsis cells functions in MAMP-elicited defence. Plant Physiol. 158,2013–2027. doi: 10.1104/pp.111.190140
Oelofse, D., Dubery, I. A., Meyer, R., Arendse, M. S., Gazendam, I., and Berger,D. K. (2006). Apple polygalacturonase inhibiting protein1 expressed in trans-genic tobacco inhibits polygalacturonases from fungal pathogens of appleand the anthracnose pathogen of lupins. Phytochemistry 67, 255–263. doi:10.1016/j.phytochem.2005.10.029
Oh, D.-H., Dassanayake, M., Bohnert, H. J., and Cheeseman, J. M. (2012). Life atthe extreme: lessons from the genome. Genome Biol. 13, 241. doi: 10.1186/gb-2012-13-3-241
Oliveira, M. B., Nascimento, L. B., Junior, M. L., and Petrofeza, S. (2010). Char-acterization of the dry bean polygalacturonase-inhibiting protein (PGIP) genefamily during Sclerotinia sclerotiorum (Sclerotiniaceae) infection. Genet. Mol.
Res. 9, 994–1004. doi: 10.4238/vol9-2gmr776Pérez-Donoso, A. G., Sun, Q., Roper, M. C., Greve, L. C., Kirkpatrick, B., and
Labavitch, J. M. (2010). Cell wall-degrading enzymes enlarge the pore size ofintervessel pit membranes in healthy and Xylella fastidiosa-infected grapevines.Plant Physiol. 152, 1748–1759. doi: 10.1104/pp.109.148791
Powell, A. L. T., Stotz, H. U., Labavitch, J. M., and Bennett, A. B. (1994). “Gly-coprotein inhibitors of fungal polygalacturonases,” in Advances in Molecular
Genetics of Plant-Microbe Interactions, Current Plant Science and Biotechnologyin Agriculture, eds M. J. Daniels, J. A. Downie, and A. E. Osbourn (Dordrecht:Springer), 399–402.
Powell, A. L. T., van Kan, J., ten Have, A., Visser, J., Greve, L. C., Ben-nett, A. B., et al. (2000). Transgenic expression of pear PGIP in tomatolimits fungal colonization. Mol. Plant Microbe Interact. 13, 942–950. doi:10.1094/MPMI.2000.13.9.942
Prabhu, S. A., Kini, K. R., Raj, S. N., Moerschbacher, B. M., and Shetty, H. S. (2012).Polygalacturonase-inhibitor proteins in pearl millet: possible involvement inresistance against downy mildew. Acta Biochim. Biophys. Sinica. 44, 415–423.doi: 10.1093/abbs/gms015
Prabhu, S. A., Singh, R., Kolkenbrock, S., Sujeeth, N., El Gueddari, N. E., Moer-schbacher, B. M., et al. (2014). Experimental and bioinformatic characterizationof recombinant polygalacturonase-inhibitor protein from pearl millet and itsinteraction with fungal polygalacturonases. J. Exp. Bot. 65, 5033–5047. doi:10.1093/jxb/eru266
Prabhu, S. A., Wagenknecht, M., Melvin, P., Gnanesh Kumar, B. S., Veena,M., Shailasree, S., et al. (2015). Immuno-affinity purification of PglPGIP1, apolygalacturonase-inhibitor protein from pearl millet: studies on its inhibi-tion of fungal polygalacturonases and role in resistance against the downymildew pathogen.Mol. Biol. Rep. doi: 10.1007/s11033-015-3850-5. [Epub aheadof print].
Protsenko, M. A., Buza, N. L., Krinitsyna, A. A., Bulantseva, E. A., andKorableva, N. P. (2008). Polygalacturonase-inhibiting protein is a struc-tural component of plant cell wall. Biochem. (Moscow) 73, 1053–1062. doi:10.1134/S0006297908100015
Raiola, A., Sella, L., Castiglioni, C., Balmas, V., and Favaron, F. (2008). A singleamino acid substitution in highly similar endo-PGs from Fusarium verticil-
lioides and related Fusarium species affects PGIP inhibition. Fungal Genet. Biol.45, 776–789. doi: 10.1016/j.fgb.2007.11.003
Reignault, P., Valette-Collet, O., and Boccara, M. (2008). The importance of fun-gal pectinolytic enzymes in plant invasion, host adaptability and symptom type.Eur. J. Plant Pathol. 120, 1–11. doi: 10.1007/s10658-007-9184-y
Richter, A., Jacobsen, H. J., De Kathen, A., De Lorenzo, G., Briviba, K., Hain, R.,et al. (2006). Transgenic peas (Pisum sativum) expressing polygalacturonaseinhibiting protein from raspberry (Rubus idaeus) and stilbene synthase fromgrape (Vitis vinifera). Plant Cell Rep. 25, 1166–1173. doi: 10.1007/s00299-006-0172-z
Frontiers in Plant Science | www.frontiersin.org 16 March 2015 | Volume 6 | Article 146
Kalunke et al. PGIP and crop protection
Ridley, B. L., O’Neill, M. A., and Mohnen, D. (2001). Pectins: structure, biosynthe-sis, and oligogalacturonide-related signaling. Phytochemistry 57, 929–967. doi:10.1016/S0031-9422(01)00113-3
Roper, M. C., Greve, L. C., Warren, J. G., Labavitch, J. M., and Kirkpatrick, B.C. (2007). Xylella fastidiosa requires polygalacturonase for colonization andpathogenicity in Vitis vinifera grapevines. Mol. Plant Microbe Interact. 20,411–419. doi: 10.1094/MPMI-20-4-0411
Sathiyaraj, G., Srinivasan, S., Subramanium, S., Kim, Y. J., Kim, Y. J., Kwon, W.S., et al. (2010). Polygalacturonase inhibiting protein: isolation, developmentalregulation and pathogen related expression in Panax ginseng C.A. Meyer. Mol.
Biol. Rep. 37, 3445–3454. doi: 10.1007/s11033-009-9936-1Sato, A., andMiura, K. (2011). Root architecture remodeling induced by phosphate
starvation. Plant Signal. Behav. 6, 1122–1126. doi: 10.4161/psb.6.8.15752Sawaki, Y., Iuchi, S., Kobayashi, Y., Kobayashi, Y., Ikka, T., Sakurai, N., et al. (2009).
STOP1 regulates multiple genes that protect Arabidopsis from proton andaluminum toxicities. Plant Physiol. 150, 281–294. doi: 10.1104/pp.108.134700
Schacht, T., Unger, C., Pich, A., andWydra, K. (2011). Endo- and exopolygalactur-onases of Ralstonia solanacearum are inhibited by polygalacturonase-inhibitingprotein (PGIP) activity in tomato stem extracts. Plant Physiol. Biochem. 49,377–387. doi: 10.1016/j.plaphy.2011.02.001
Sella, L., Castiglioni, C., Roberti, S., D’Ovidio, R., and Favaron, F. (2004). Anendo-polygalacturonase (PG) of Fusarium moniliforme escaping inhibitionby plant polygalacturonase-inhibiting proteins (PGIPs) provides new insightsinto the PG-PGIP interaction. FEMS Microbiol. Lett. 240, 117–124. doi:10.1016/j.femsle.2004.09.019
Senthil, G., Williamson, B., Dinkins, R. D., and Ramsay, G. (2004). An efficienttransformation system for chickpea (Cicer arietinum L.). Plant Cell Rep. 23,297–303. doi: 10.1007/s00299-004-0854-3
Shanmugam, V. (2005). Role of extracytoplasmic leucine rich repeat pro-teins in plant defence mechanisms. Microbiol. Res. 160, 83–94. doi:10.1016/j.micres.2004.09.014
Shivashankar, S., Thimmareddy, C., and Roy, T. K. (2010). Polygalacturonaseinhibitor protein from fruits of anthracnose resistant and susceptible varietiesof Chilli (Capsicum annuum L). Indian J. Biochem. Biophys. 47, 243–248.
Sicilia, F., Fernandez-Recio, J., Caprari, C., De Lorenzo, G., Tsernoglou, D.,Cervone, F., et al. (2005). The polygalacturonase-inhibiting protein PGIP2of Phaseolus vulgaris has evolved a mixed mode of inhibition of endopoly-galacturonase PG1 of Botrytis cinerea. Plant Physiol. 139, 1380–1388. doi:10.1104/pp.105.067546
Song, K. H., and Nam, Y. W. (2005). Genomic organization and differentialexpression of two polygalacturonase-inhibiting protein genes from Medicago
truncatula. J. Plant Biol. 48, 467–478. doi: 10.1007/BF03030589Spadoni, S., Zabotina, O., Di Matteo, A., Mikkelsen, J. D., Cervone, F., De Lorenzo,
G., et al. (2006). Polygalacturonase-inhibiting protein interacts with pectinthrough a binding site formed by four clustered residues of arginine and lysine.Plant Physiol. 141, 557–564. doi: 10.1104/pp.106.076950
Spinelli, F., Mariotti, L., Mattei, B., Salvi, G., Cervone, F., and Caprari, C. (2009).Three aspartic acid residues of polygalacturonase-inhibiting protein (PGIP)from Phaseolus vulgaris are critical for inhibition of Fusarium phyllophilum PG.Plant Biol. 11, 738–743. doi: 10.1111/j.1438-8677.2008.00175.x
Szankowski, I., Briviba, K., Fleschhut, J., Schönherr, J., Jacobsen, H. J., andKiesecker, H. (2003). Transformation of apple (Malus domestica Borkh.)with the stilbene synthase gene from grapevine (Vitis vinifera L.) and aPGIP gene from kiwi (Actinidia deliciosa). Plant Cell Rep. 22, 141–149. doi:10.1007/s00299-003-0668-8
Tamburino, R., Chambery, A., Parente, A., and Di Maro, A. (2012). A NovelPolygalacturonase-Inhibiting Protein (PGIP) from Lathyrus sativus L. seeds.Protein Pept. Lett. 19, 820–825. doi: 10.2174/092986612801619561
Tamura, M., Gao, M., Tao, R., Labavitch, J. M., and Dandekar, A. M. (2004).Transformation of persimmon with a pear fruit polygalacturonase inhibit-ing protein (PGIP) gene. Sci. Hortic. 103, 19–30. doi: 10.1016/j.scienta.2004.04.006
ten Have, A., Mulder, W., Visser, J., and van Kan, J. A. (1998). The endopolygalac-turonase gene Bcpg1 is required for full virulence of Botrytis cinerea.Mol. Plant
Microbe Interact. 11, 1009–1016. doi: 10.1094/MPMI.1998.11.10.1009Thornburg, R. W., Carter, C., Powell, A. L., Mittler, R., Rizhsky, R., and Horner,
H. T. (2003). A major function of the tobacco floral nectary is defense againstmicrobial attack. Plant Syst. Evol. 238, 211–218. doi: 10.1007/s00606-003-0282-9
Toubart, P., Desiderio, A., Salvi, G., Cervone, F., Daroda, L., De Lorenzo,G., et al. (1992). Cloning and characterization of the gene encoding theendopolygalacturonase-inhibiting protein (PGIP) of Phaseolus vulgaris L. PlantJ. 2, 367–373. doi: 10.1046/j.1365-313X.1992.t01-35-00999.x
van Santen, Y., Benen, J. A., Schroter, K. H., Kalk, K. H., Armand, S.,and Visser, J. (1999). 1.68-Å crystal structure of endopolygalacturonase IIfrom Aspergillus niger and identification of active site residues by site-directed mutagenesis. J. Biol. Chem. 274, 30474–30480. doi: 10.1074/jbc.274.43.30474
Volpi, C., Raiola, A., Janni, M., Gordon, A., O’Sullivan, D. M., Favaron, F.,et al. (2013). Claviceps purpurea expressing polygalacturonases escaping PGIPinhibition fully infects PvPGIP2 wheat transgenic plants but its infectionis delayed in wheat transgenic plants with increased level of pectin methylesterification. Plant Physiol. Biochem. 73, 294–301. doi: 10.1016/j.plaphy.2013.10.011
Wand, A. J., and Englander, S. W. (1996). Protein complexes studied by NMRspectroscopy. Curr. Opin. Biotechnol. 7, 403–408.
Wang, A., Wei, X., Rong, W., Dang, L., Du, L. P., Qi, L., et al. (2014a). GmPGIP3enhanced resistance to both take-all and common root rot diseases in trans-genic wheat. Funct. Integr. Genomics. doi: 10.1007/s10142-014-0428-6. [Epubahead of print].
Wang, R., Lu, L., Pan, X., Hu, Z., Ling, F., Yan, Y., et al. (2014b). Functional analysisof OsPGIP1 in rice sheath blight resistance. Plant Mol. Biol. 87, 181–191. doi:10.1007/s11103-014-0269-7
Wang, X., Zhu, X., Tooley, P., and Zhang, X. (2013). Cloning and func-tional analysis of three genes encoding polygalacturonase-inhibiting proteinsfrom Capsicum annuum and transgenic CaPGIP1 in tobacco in relation toincreased resistance to two fungal pathogens. Plant Mol. Biol. 81, 379–400. doi:10.1007/s11103-013-0007-6
Conflict of Interest Statement: The authors declare that the research was con-ducted in the absence of any commercial or financial relationships that could beconstrued as a potential conflict of interest.
Copyright © 2015 Kalunke, Tundo, Benedetti, Cervone, De Lorenzo and D’Ovidio.
This is an open-access article distributed under the terms of the Creative Commons
Attribution License (CC BY). The use, distribution or reproduction in other forums
is permitted, provided the original author(s) or licensor are credited and that the
original publication in this journal is cited, in accordance with accepted academic
practice. No use, distribution or reproduction is permitted which does not comply
with these terms.
Frontiers in Plant Science | www.frontiersin.org 17 March 2015 | Volume 6 | Article 146
152
Superhydrophobicity in perfection:the outstanding properties of the lotus leafHans J. Ensikat*1, Petra Ditsche-Kuru1, Christoph Neinhuis2
and Wilhelm Barthlott1
Full Research Paper Open Access
Address:1Nees Institute, University of Bonn, Meckenheimer Allee 170, 53115Bonn, Germany and 2Institut für Botanik, Technische UniversitätDresden, Zellescher Weg 20b, 01069 Dresden, Germany
Email:Hans J. Ensikat* - [email protected]
* Corresponding author
Keywords:epicuticular wax; leaf surface; Lotus effect; papillae; water repellency
Beilstein J. Nanotechnol. 2011, 2, 152–161.doi:10.3762/bjnano.2.19
Received: 07 January 2011Accepted: 17 February 2011Published: 10 March 2011
This article is part of the Thematic Series "Biomimetic materials".
Guest Editors: W. Barthlott and K. Koch
© 2011 Ensikat et al; licensee Beilstein-Institut.License and terms: see end of document.
AbstractLotus leaves have become an icon for superhydrophobicity and self-cleaning surfaces, and have led to the concept of the ‘Lotus
effect’. Although many other plants have superhydrophobic surfaces with almost similar contact angles, the lotus shows better
stability and perfection of its water repellency. Here, we compare the relevant properties such as the micro- and nano-structure, the
chemical composition of the waxes and the mechanical properties of lotus with its competitors. It soon becomes obvious that the
upper epidermis of the lotus leaf has developed some unrivaled optimizations. The extraordinary shape and the density of the
papillae are the basis for the extremely reduced contact area between surface and water drops. The exceptional dense layer of very
small epicuticular wax tubules is a result of their unique chemical composition. The mechanical robustness of the papillae and the
wax tubules reduce damage and are the basis for the perfection and durability of the water repellency. A reason for the optimiza-
tion, particularly of the upper side of the lotus leaf, can be deduced from the fact that the stomata are located in the upper epidermis.
Here, the impact of rain and contamination is higher than on the lower epidermis. The lotus plant has successfully developed an
excellent protection for this delicate epistomatic surface of its leaves.
152
IntroductionSince the introduction of the ‘Lotus concept’ in 1992 [1,2], the
lotus leaf became the archetype for superhydrophobicity and
self-cleaning properties of plant surfaces and a model for tech-
nical analogues [3,4] . Lotus (Nelumbo nucifera) is a semi-
aquatic plant and develops peltate leaves up to 30 cm in dia-
meter with remarkable water repellency. As an adaptation to the
aquatic environment – some of the leaves float occasionally on
the water surface – the stomata are located in the upper
Beilstein J. Nanotechnol. 2011, 2, 152–161.
153
epidermis. The lower epidermis consists of convex cells
covered with wax tubules and contains only few stomata. The
upper epidermis features the distinctive hierarchical structure
consisting of papillae with a dense coating of agglomerated wax
tubules, which is the basis for the famous superhydrophobicity
(Figure 1).
Figure 1: (a) Lotus leaves, which exhibit extraordinary water repel-lency on their upper side. (b) Scanning electron microscopy (SEM)image of the upper leaf side prepared by ‘glycerol substitution’ showsthe hierarchical surface structure consisting of papillae, wax clustersand wax tubules. (c) Wax tubules on the upper leaf side. (d) Upper leafside after critical-point (CP) drying. The wax tubules are dissolved,thus the stomata are more visible. Tilt angle 15°. (e) Leaf underside(CP dried) shows convex cells without stomata.
However, a hierarchical surface structure which induces strong
water repellency and contact angles above 150° is not a special
feature of lotus leaves. It has been known for a long time that
plant surfaces covered with epicuticular wax crystals are water
repellent, and that this feature is enhanced when the epidermis
has additional structures such as papillae or hairs [5,6]. Nein-
huis and Barthlott (1997) [7] presented an overview of more
than 200 species with contact angles >150° and their surface
morphologies. Many studies, in which the properties of lotus
leaves were compared with those of other superhydrophobic
plants, have shown the superiority of the upper side of the lotus
leaf. A standard tool for the determination of wettability or
water repellency is the measurement of the static contact angle
by the ‘sessile drop’ method. Neinhuis and Barthlott (1997) [7]
for example, measured contact angles on the lotus leaf of 162°,
which are among the highest of the compared species, but many
other (43%) of the tested superhydrophobic plants also showed
contact angles between 160 and 163°. Even some species with
flat epidermis cells but with a dense layer of epicuticular wax
crystals, such as Brassica oleracea or some Eucalyptus species,
can exhibit contact angles >160°. Thus, the contact angle alone
is not suitable for a differentiated comparison of superhy-
drophobic samples. Other values such as contact angle
hysteresis or roll-off (tilting) angle show more clearly the
differences between the species. Mockenhaupt et al. (2008) [8]
compared the tilting angles and the stability of the superhy-
drophobicity of various plants under moisture condensation
conditions. Only the lotus leaves showed no significant loss of
water repellency when water vapour condensed on the surface
of the cooled samples at 5 °C. Wagner et al. (2003) [9] exam-
ined the morphology of the epidermal structures and the wetta-
bility with liquids of varying surface tension such as
methanol–water mixtures. They reported the lowest wettability
by these liquids for the lotus leaves in comparison to other
species. They also described the unique shape of the papillae
and a very high papillae density (number per area). Chemical
analyses [10] and crystal structure analysis by X-ray diffraction
[11] showed unique properties of the epicuticular wax of the
lotus. The high content of nonacosanediols leads to a high
melting point as well as a strongly disturbed crystal structure
which is the basis for the formation of tubules. The visualiza-
tion of the contact zone between leaves and droplets with cryo-
scanning electron microscopy demonstrated the extremely
reduced contact area for lotus [12]. Zhang et al. (2008) [13]
made detailed measurements of the water repellency of the
papillose lotus leaf surface in comparison with the non-papil-
lose leaf margin. The importance of the nanoscopic wax crys-
tals for the water repellency was demonstrated by Cheng et al.
(2006) [14]. They reported a strong decrease of the contact
angle after melting of the waxes. A limited air retaining capa-
bility of submersed lotus leaves was reported by Zhang et al.
(2009) [15] after the leaves were held at a depth of 50 cm for
2 h. Bhushan et al. (2010) [4] used the surface structures of the
lotus leaf as model for the development of artificial biomimetic
superhydrophobic structures.
It became obvious that the outstanding and stable superhy-
drophobicity of the lotus leaf relies on the combination of opti-
mized features such as the surface topography, robustness and
the unique properties of the epicuticular wax. The aim of this
article is to integrate the relevant features of the lotus leaf, and
to compare them with superhydrophobic leaves of other plant
species in order to illustrate their significance.
Beilstein J. Nanotechnol. 2011, 2, 152–161.
154
Figure 2: Epidermis cells of the leaf upper side with papillae. The surface is densely covered with wax tubules. (a) SEM image after freeze drying. (b)Light microscopy (LM) image of thin section of an embedded sample. Assuming a contact angle of >140°, for example, the area of heterogeneouscontact between single papillae and water (marks) is small in comparison to the epidermis cell area.
Results and DiscussionThe properties of the lotus leavesThe lotus leaf shows an outstanding water repellency particu-
larly on its upper (adaxial) side, which is more robust and less
sensitive to mechanical damage than the under (abaxial) side.
The reasons for these superior properties can be ascribed to the
combination of micro- and nano-structures with optimized
geometry and the unique chemical composition of the epicutic-
ular waxes. These properties are illustrated in the following
sections and compared with those of other superhydrophobic
leaves. (The species are listed in the Experimental section).
Minimization of the water-to-leaf contact area: The
epidermis cells of the upper leaf side form papillae of varying
height and with a unique shape. The diameter of the papillae is
much smaller than that of the epidermis cells and each papilla
apex is not spherical but forms an ogive (Figure 2).
The whole surface is covered with short wax tubules which
often accumulate in clusters. In comparison with other papil-
lose plant surfaces, lotus has the highest density of papillae, but
the lotus papillae have much smaller diameters which reduces
the contact area with water drops; strictly speaking, the area of
heterogeneous contact between surface and water. The contact
area depends on the hydrophobicity of the surface and on the
pressure of the water or on the kinetic energy or velocity of the
striking water drops. At low pressures, caused by resting or
rolling water droplets, the contact area is determined by the
local contact angle of the surface structures. For the surface of a
papilla coated with wax tubules, a superhydrophobic behavior
with a local contact angle of >140° can be assumed. So, the dia-
meter of the contact areas can be estimated from the SEM
images and the cross sections of the selected samples
(Figure 2).
The minimized contact area is the basic cause for the very low
adhesion of water and, thus, the small roll-off (tilting) angles.
Compared with lotus, the papillae on the leaves of the other
plants (E. myrsinites, C. esculenta, A. macrorrhiza) (Figure 3,
see also Figure 7) have much larger diameters and tip radii, and
are covered with different wax types, wax platelets or wax film,
respectively, which have a lower water repellency than wax
tubules.
Figure 3: SEM images of the papillose leaf surfaces of Nelumbonucifera (Lotus) (a), Euphorbia myrsinites (b), Colocasia esculenta (c),and Alocasia macrorrhiza (d). Lotus has the highest density of papillaewith varying heights and the smallest diameter of the papillae. Thepapillae of the other species have larger diameters and are coveredwith different wax types: wax platelets (E. myrsinites and C. esculenta)and a wax film (A. macrorrhiza) which covers cuticular foldings.
Beilstein J. Nanotechnol. 2011, 2, 152–161.
155
Figure 4: The contact between water and superhydrophobic papillaeat different pressures. At moderate pressures the water intrudes intothe space between the papillae, but an air layer remains betweenwater and epidermis cells (a). The superhydrophobic surface of thepapillae causes a repellent force (‘re’). When the water recedes, thenthe papillae lose contact one after the other (b, c). At a certain waterlevel, the meniscus is flat and the force is neutral (‘n’). Just prior to theseparation an adhesive force (‘ad’) arises at the almost horizontal areaof the papilla tip, which is small on tips with intact wax crystals andlarger when the wax is damaged or eroded. On artificial superhy-drophobic structures with equal height (d) the adhesive forces duringwater receding occur simultaneously at all contacts.
The varying height of the papillae further reduces the adhesion
between water drops and the surface (Figure 4). Small resting or
sliding water drops touch only the highest papillae [12]. At
higher pressures, e.g., at the impact of raindrops, the water
intrudes deeper between the papillae (Figure 4a) and forms a
meniscus at the still superhydrophobic wax tubules coating. The
deformation of the non-wetting droplet surface due to surface
tension causes a repellent force (‘re’, Figure 4). When the water
retracts, either at the receding side of a moving drop or if the
drop is lifted off the surface, the contact areas decrease and the
papillae release their contact to the water one by one (Figure 4b,
Figure 4c), so that only few of the papillae are simultaneously
in the adhesive state (‘ad’). Finally, before the drop loses
contact with the leaf, only few of the papillae are still in contact
and cause a small adhesive force. In contrast, artificial superhy-
drophobic samples with pillars of equal height lead to stronger
adhesion during drop retraction when all the pillars are simulta-
neously in the adhesive state before the contact breaks
(Figure 4d). The measurement of the adhesive and repellent
forces between a superhydrophobic papilla-model (with ten
times larger tip radius than a lotus papilla) and a water drop is
shown in Figure 5.
Figure 5: Measured forces between a superhydrophobic papilla-modeland a water drop during advancing and receding. The images corres-ponding to the marks (arrows) in the diagram show the repellent (a)and adhesive (b) meniscus. (c) Papilla-model tip shown with SEM.
Contact angle measurements are the standard tool for the
determination of hydrophobicity. But the measurement of very
high contact angles is often inaccurate due to difficulties in the
determination of the exact drop shape [16], particularly on
uneven leaf surfaces. For many superhydrophobic plant
surfaces, the contact angles are very close together [7] such that
the inaccuracies are larger than the differences between the
samples. This may prevent a meaningful comparison. A more
differentiated comparison of water repellency has been
achieved by the measurement of the adhesion between surface
and water during retraction of a drop [13], similar to the
measurement shown in Figure 5. Table 1 shows, in addition to
other relevant properties, the maximal adhesion forces of water
drops on fresh lotus leaves and leaves of other species with
intact wax. The adhesion forces are strongly dependent on
Beilstein J. Nanotechnol. 2011, 2, 152–161.
156
Table 1: Comparison of water repellency relevant properties of lotus and other selected species.
Nelumbo nucifera(Lotus)(upper side)
Colocasiaesculenta(upper side)
Euphorbiamyrsinites(upper side)
Alocasiamacrorrhiza(lower side)
Brassicaoleracea(upper side)
papillae density (per mm2) [9] 3431 2662 1265 2002 0contact angle (static) [7] 163° 165° 162° 157° 161°drop adhesion force (µN)a 8–18 28–55 30–58 90–127 7–48
wax type tubules platelets platelets film on cuticularfolds
rodlets andtubules
wax melting point (°C) 90–95 75–78 75–76 n.a.b 65–67
main components [11] C29-diols C28-1-ol C26-1-ol n.a.b C29-ketones,C29-alkanes
aprovided by D. Mohr, Nees Institute, Bonn; bthe wax film of A. macrorrhiza has not been isolated and analyzed; no data available.
Figure 6: Papillose and non-papillose leaf surfaces with an intactcoating of wax crystals: (a) Nelumbo nucifera (Lotus); (b) Euphorbiamyrsinites; (c) Brassica oleracea; (d) Yucca filamentosa. Even thenon-papillose leaves are superhydrophobic. The contact angle of B.oleracea can exceed 160°.
surface defects which cause pinning of the drops. In contrast,
advancing contact angles depend weakly on such irregularities.
Thus, the adhesion data correlate better with receding contact
angles and hysteresis and indicate the perfection and defects of
superhydrophobic surfaces.
Mechanical protection of the wax crystals by papillae: The
highest water repellency occurs when the water drops touch the
tips of the epicuticular wax crystals only. Thus, the best prop-
erties are found on leaves with an intact coating of wax crystals
on the epidermal cells (Figure 6). The waxes are, however, rela-
tively soft materials so that older leaves often show patches of
eroded or damaged wax (Figure 7), which cause an increased
adhesion of water. Neinhuis and Barthlott (1997) [7] have
reported that papillae protect the wax crystals between them. On
papillose epidermis cells only the wax on the papillae tips
Figure 7: Traces of natural erosion of the waxes on the same leavesas in Figure 6: (a) Nelumbo nucifera (Lotus); (b) Euphorbia myrsinites;(c) Brassica oleracea; (d) Yucca filamentosa. On the papillose leaves(a,b) the eroded areas are limited to the tips of the papillae. On non-papillose cells, the damaged areas can be much larger (c,d), causingstronger pinning of water droplets.
appears damaged while the wax between the papillae remains
intact (Figure 7a, Figure 7b). Thus, lotus leaves retain their
water repellency up to the end of their lifetime. In contrast, the
non-papillose surfaces of Brassica oleracea and Yucca filamen-
tosa (Figure 7c, Figure 7d) often show larger damaged areas
which cause a stronger pinning of water. The efficiency of the
protective properties can easily be tested by wiping across the
leaf with the finger, which destroys only the wax on the papillae
tips (Figure 8a, Figure 8b), but the leaves remained superhy-
drophobic. In the case of the non-papillose surface of a B. oler-
acea leaf (Figure 8c), the waxes are completely destroyed and
superhydrophobicity is lost; the contact angle decreased from
160° to ca. 130°. On a Y. filamentosa leaf (Figure 8d) with
convex epidermis cells, most of the wax crystals were destroyed
and the contact angle dropped from 150° to ca. 110°.
Beilstein J. Nanotechnol. 2011, 2, 152–161.
157
Figure 8: Test for the stability of the waxes against damaging bywiping on the same leaves: (a) Nelumbo nucifera (Lotus); (b)Euphorbia myrsinites; (c) Brassica oleracea; (d) Yucca filamentosa. Onthe papillose surfaces only the waxes on the tips of the papillae aredestroyed. The waxes between the papillae are protected and remainintact. On the non-papillose surfaces, most of the waxes aredestroyed, adhesion of water drops (pinning) is strongly increased, andthe superhydrophobicity is lost.
Figure 9: SEM and LM images of cross sections through the papillae.Lotus (a,b) and Euphorbia myrsinites (c,d) have almost massivepapillae, those of Alocasia macrorrhiza (e,f) have a relatively thickouter wall; the epidermal cells of Colocasia esculenta have thin walls(g,h). The arrow in (b) marks a stoma.
The basis for the ability to protect the leaf surface in lotus is the
robustness of its leaf papillae in combination with their high
density. Cross sections (Figure 9) show that they are almost
massive at least in the apical part, in contrast to the fragile
papillose cells found on many flower petals. However, papillae
of other superhydrophobic leaves show various architectures:
Euphorbia myrsinites has completely massive papillae; those of
the lower epidermis of Alocasia macrorrhiza have quite thick
outer walls, whereas the epidermal cells of Colocasia esculenta
have very thin walls with slight thickening at the protrusions.
Properties of the lotus waxBoth the upper side and the lower side of the lotus leaf are
covered with wax tubules. But, as can be seen on the SEM
images (Figure 10a, Figure 10b), the waxes of both sides look
quite different. The wax tubules of the lower side are longer (1
to 2 μm) and thicker (ca. 150 nm) and are typical ‘nonacosanol
tubules’ which commonly occur on many plant species [7]. In
contrast, the wax tubules of the upper leaf side are very short
(0.3–1 µm) and thin (80–120 nm) but the density is very high.
Figure 10 shows on a clearly arranged area, approximately 200
tubules per 10 µm2 on the upper side, but only about 63 tubules
per 10 µm2 on the lower side of the same leaf. The spacing
between the tubules on the upper side of the lotus leaf is much
smaller than that of other wax crystals such as platelets
(Figure 10c, Figure 10d) and other tubular waxes (Figure 10b,
Figure 10e, Figure 10f). These distances between the
hydrophobic wax crystals determine the pressure (capillary
pressure) which is necessary for an intrusion of a water droplet
between them.
The chemical analyses of the waxes give an explanation for the
different properties. It is known that the epicuticular wax of
lotus contains a high percentage of nonacosanediols [10], but
the older analyses were made from the entire wax of the leaves,
which was obtained as a chloroform extract and also contained
intracuticular lipids. The new analyses of the separately isolated
waxes from both sides (Figure 11) show that the wax of the
upper side contains ca. 65% of various nonacosanediols and
only 22% of nonacosan-10-ol, whereas the wax of the under-
side contains predominantly nonacosan-10-ol (53%) and only
15% of diols, together with 18% of alkanes. The remaining 13%
and 14% could not be identified.
This high content of nonacosanediols provides extraordinary
properties to the upper side wax. The melting point of 90 to
95 °C is very high for normal (aliphatic) waxes and indicates
the influence of hydrogen bonding in the crystal lattice which
increases the stability. A comparison of different aliphatic wax
components with similar chain length shows that the melting
points increase with the occurrence of polar OH-groups. Strong
Beilstein J. Nanotechnol. 2011, 2, 152–161.
158
Figure 10: Epicuticular wax crystals in an area of 4 × 3 µm2. Theupper side of the lotus leaf (a) has the highest crystal density (numberper area) of wax crystals and the smallest spacings between them.Lotus upper side (a) ca. 200 tubules per 10 µm2; (b) Lotus undersideca. 63 tubules per 10 µm2; (c) Euphorbia myrsinites ca. 50 plateletsper 10 µm2; (d) Yucca filamentosa ca. 17 platelets with over 80 jagsper 10 µm2; (e) Brassica oleracea ca. 22 rodlets and tubules, and (f)Eucalyptus macrocarpa ca. 50 tubules per 10 µm2. The larger spacingbetween the wax crystals of the other surfaces compared to the lotusupper side is obvious.
hydrogen bonding effects have been measured recently by
Coward (2010) [17] in nonacosanol wax using FTIR spec-
troscopy. The effects on the crystal structure should be even
stronger for the nonacosanediols. Although the secondary alco-
hols (nonacosan-10-ol and nonacosanediols) contain polar
OH-groups in their molecules, the resulting wax tubules are
known to feature strong and relatively stable water repellency,
particularly the diols of the lotus leaf. This seems paradoxical,
but X-ray diffraction analyses (Figure 12) are in accordance
with a layer structure model in which the OH-groups are buried
deep in the layer, while the layer surface consists only of non-
polar methyl groups [11,18]. In contrast, primary alcohols such
as the widespread octacosan-1-ol, which occurs in many
platelet-shaped epicuticular waxes, can present the OH-group
on the surface, e.g., if they are in contact with a polar environ-
ment (water). Holloway (1969) [19] studied the hydrophobicity
and water contact angles of various plant waxes and pure wax
components. He found the highest contact angles for aliphatic
waxes which present only methyl groups on the surface.
According to the layer structure model, the tubules are strongly
curved helically growing layers. While straight long-chained al-
Figure 11: Chemical composition of the separated waxes of the upperand lower side of the lotus leaf. The upper side wax contains 65% ofvarious diols and only 22% of nonacosan-10-ol (C29-10-ol), 13% wasunidentified; the underside wax contains 53% nonacosan-10-ol andonly 15% of various diols. Alkanes (18%) were only found in the under-side wax and may be an essential part of the underlying wax film.
Figure 12: X-ray diffraction diagram of upperside lotus wax. The ‘longspacing’ peaks indicate a layer structure which is common in aliphaticwaxes. The broad ‘short spacing’ peak at 2θ = 27° indicates a strongdisorder in the lateral package of the molecules.
kane molecules form flat layers and regular platelet crystals,
secondary alcohols and ketones carry lateral oxygen atoms
which inhibit a tight package of the molecules. Thus the
resulting layers have a strong curvature and form tubules with a
circular cross-section (Figure 13). Today, the progress in molec-
Beilstein J. Nanotechnol. 2011, 2, 152–161.
159
Figure 13: Model of a wax tubule composed of layers of nonacosan-10-ol and nonacosanediol molecules. The OH-groups (red) occupyadditional space so that the dense package is disturbed and the layeris forced into a curvature which leads to the formation of a tubule. Thepolar OH-groups are hidden in the layer, only the CH3-groups appearat the surface of the layers and tubules.
ular dynamics simulations enables the calculation of the behav-
iour of nano-structured surfaces in contact with water [20] and
to prove the theories such as those of Wenzel or Cassie and
Baxter. For precise modelling of the behaviour of natural water
repellent surfaces, an exact knowledge of the chemical compos-
ition and molecular structure are essential.
Resistance against environmental stressThe excellent superhydrophobic properties of the upper side of
the lotus leaf are a result of several unique optimizations. The
question then arises whether this development has a certain
reason or whether it is a ‘freak of nature’. On most plants, the
undersides of the leaves show the highest water repellency, or
more precisely, those sides which are equipped with stomata. It
is obvious that the water repellency serves as a protection to
keep the stomata dry [7]. On some species only the cells around
the stomata are covered with wax crystals. This is in accor-
dance with the fact that the lotus leaf is epistomatic; it bears the
stomata on the upper side, which possesses the higher water
repellency. The upper side of a leaf is strongly exposed to envi-
ronmental impacts such as rainfall and deposition of contamina-
tions. Obviously it is a greater challenge to keep the upper side
of a large leaf dry and clean than the underside or the surfaces
of vertically growing leaves (grasses etc.). On most plants, the
upper sides of the leaves bear no stomata and are more robust
than the undersides [21]. Thus the extremely stable and durable
water repellency of the lotus leaf, which persists up to the end
of its lifetime in autumn, seems to be a successful evolutionary
adaptation to the aquatic environment, which led to the placing
of the stomata in the upper epidermis and the development of an
effective protection through specialized epidermal structures.
For a stable superhydrophobicity – that means the retention of
the Cassie state with only partial contact between surface and
water – an intrusion of water between the surface structures
must be avoided. When the air layer is displaced by water, the
water repellency is lost and the surface becomes wet (Wenzel
state). The pressure which is necessary to press water into the
space between hydrophobic structures depends on the local
contact angle and the size of the spacing. This pressure (capil-
lary pressure) is reciprocal to the size of the spacing and can be
deduced from the Young–Laplace equation. Due to the irreg-
ular spacing, it can be estimated roughly. Water droplets with a
radius <100 nm may be able to intrude between the wax
tubules; this curvature corresponds to a Laplace pressure of
>1.4 MPa (14 bar). Varanasi et al. (2009) [22] calculated the
capillary pressures of hydrophobic test samples with structure
dimensions roughly similar to those of the lotus leaf: The capil-
lary pressure for spacing of 5 µm between hydrophobic pillars
is 12 kPa (120 mbar); a nanoporous structure with 90 nm pore
diameter has a capillary pressure of 1.6 MPa (16 bar). Thus the
capillary pressure of the lotus papillae with spacing of ca.
10 µm is sufficient to carry the load of resting or rolling water
drops. But impacting raindrops generate higher pressure pulses
and can intrude into the space between the papillae. The
maximal pressure for a drop impact on a rigid material can be
calculated from the ‘water hammer’ equation: pWH = 0.2 ρ·c·v,
where ρ is the density of the liquid, c is the speed of sound in
the liquid, and v is the velocity of the droplet. Varanasi et al.
(2009) [22] calculated the ‘water hammer pressure’ of rain-
drops with a velocity of 3 m/s as 0.9 MPa (9 bar). However,
drop impacts on flexible surfaces generate considerably lower
pressures [23]. Due to the small spacing between the wax
tubules of the lotus leaf and their strong hydrophobicity, their
capillary pressure is obviously higher than the impact pressure
of raindrops and sufficient to prevent water intrusion. However,
it is unproven and hypothetical whether the larger spacing in
other waxes causes an intrusion of raindrops. Mechanical
damage to the waxes by the impacting drops is a more likely
cause for degradation.
Biological models serve as an inspiration for the development
of technical superhydrophobic materials [4]. So the question
arises whether the lotus leaf presents an optimal architecture for
superhydrophobicity. In biological surfaces, several different
strategies can be found. The lotus leaf with the largely reduced
contact area seems optimal for low adhesion of contaminants
and water, observable as small roll-off angles. A disadvantage
is the relatively soft wax material, which is too fragile for most
technical applications. A different architecture is found on some
species with hairy leaf surfaces. The water fern (some species
of the genus Salvinia) and Pistia stratioides leaves retain a rela-
tively thick air layer between hydrophobic hairs when sub-
Beilstein J. Nanotechnol. 2011, 2, 152–161.
160
mersed in water [24]. This provides sufficient buoyancy to
avoid long-term submerging. Although superhydrophobic
leaves retain an air layer when they are submersed, they are not
designed for continuously living under water. All permanently
submersed plant surfaces are hydrophilic without hydrophobic
waxes [25]. Superhydrophobic surfaces which feature perma-
nent air retention under water are found on animals (some birds,
spiders and insects). An outstanding air-retention capability is
found, for example, for the aquatic insect Notonecta glauca
(‘backswimmer’) [26,27]. Here the water repellency is created
by a two-level structure consisting of coarse hairs which can
hold a relatively thick air layer, and extremely fine hairs which
ensure a high capillary pressure. The biopolymers used in these
structures have the advantage of a much higher strength than
waxes. On the other hand, the plant surfaces have the capability
to regenerate damaged or lost waxes.
ConclusionIt is true that lotus exhibits outstanding water repellency on the
upper side of its leaves. The basis of this behaviour is the hier-
archical surface structure. In comparison to other species with a
hierarchical surface structure composed of papillae and wax
crystals, the lotus leaf shows special optimization of some of its
features. The morphology of the papillae, particularly the small
tip radius, minimizes the contact area to water drops but also the
area where erosion and damaging of the waxes occurs. The
robustness of the papillae ensures protection of the wax crystals
between them. The chemical composition of the epicuticular
wax with the high content of nonacosanediols leads to the
growth of a dense layer of very small wax tubules with a perma-
nently hydrophobic surface. The unique combination of these
properties provides the lotus leaves with unrivaled superhy-
drophobicity and self-cleaning properties as an effective protec-
tion of the delicate epistomatic surface.
ExperimentalIn addition to the data from the literature, some new examina-
tions provided material for this publication. Plant leaves were
taken from the Botanical Gardens, University of Bonn: Alocasia
macrorrhiza (Elephant ear), Brassica oleracea var. gongylodes
(Kohlrabi), Colocasia esculenta (Taro), Euphorbia myrsinites,
Nelumbo nucifera (Lotus), Yucca filamentosa.
For scanning electron microscopy, a Cambridge Stereoscan
S200 SEM was used. Depending on the sample properties,
different preparation methods were applied: Slowly drying
leaves were examined as fresh-hydrated samples (Euphorbia
myrsinites, Alocasia macrorrhiza, Brassica oleracea, Yucca
filamentosa). The other species were critical-point dried or
freeze dried (Lotus). Air-dried samples were used for high-
magnification imaging of epicuticular waxes. These prepara-
tion methods are described in detail elsewhere [28]. The
samples for thin sections were prepared following a standard
protocol for transmission electron microscopy preparation [29]:
fixation in glutaraldehyde, dehydration with acetone, embed-
ding in epoxy resin (Agar Low Viscosity Kit, Plano GmbH,
Wetzlar, Germany). Sections of ca. 0.5 µm thickness were
stained with ‘Rapid dye’ (Azur II and Methylene blue) for light
microscopy.
Wax samples for chemical analyses were isolated mechanically
using a ‘cryo-adhesion’-method using triethylene glycol as
preparation liquid [30]. The wax was analysed by gas chroma-
tography (HP 5890 series II, Avondale, USA) after ‘derivatiza-
tion’ by the reaction with N,O-bis(trimethylsilyl)trifluoroacet-
amide [31]. X-ray powder diffraction diagrams were recorded
with a diffractometer PW 1049/10 (Philips, Eindhoven, The
Netherlands) [6].
Contact angles of water drops on the sample surfaces were
measured with a contact angle measurement system (OCA 30-2,
Dataphysics Instruments GmbH, Filderstadt, Germany) using
drops of 10 µL. The adhesion of water drops on the samples
was measured with a self-developed device by recording
force–distance curves while the drop was attached to and
detached from the surface with constant velocity. Drops of
10 µL with a diameter of 2.5 mm were attached until the contact
area was 0.7 mm in diameter. Then the maximal adhesion
forces during retraction were measured and compared. Low
adhesion forces correlate with strong water repellency. The
robustness of the leaf surface structures was tested by wiping
the leaves with a finger, with a vertical force of 1 N and a
contact area of 2.5 cm2.
AcknowledgementsOur studies were supported by the Bundesministerium für
Bildung, Forschung und Technologie (BMBF), the German
Science Foundation (Deutsche Forschungsgemeinschaft, DFG)
and the Akademie der Wissenschaften und der Literatur zu
Mainz (Project V ’Biodiversität im Wandel’). We thank Prof.
Lukas Schreiber, Bonn, for the chemical analyses, Prof. Kerstin
Koch (Kleve) and Ms. Divykriti Chopra for the help in the prep-
aration of the manuscript, and Dominic Mohr (Bonn) for assis-
tance with the measurements.
References1. Barthlott, W. Die Selbstreinigungsfähigkeit pflanzlicher Oberflächen
durch Epicuticularwachse. In Klima- und Umweltforschung an derUniversität Bonn; Rheinische Friedrich-Wilhelms-Universität, Bonn,Ed.; Bornemann: Bonn, 1992; pp 117–120.
2. Barthlott, W.; Neinhuis, C. Planta 1997, 202, 1–8.doi:10.1007/s004250050096
Beilstein J. Nanotechnol. 2011, 2, 152–161.
161
3. Barthlott, W.; Neinhuis, C. Biol. Unserer Zeit 1998, 28, 314–321.doi:10.1002/biuz.960280507
4. Bhushan, B.; Jung, C. J.; Nosonovsky, M. Lotus Effect: Surfaces withRoughness-Induced Superhydrophobicity, Self-Cleaning, and LowAdhesion. In Springer Handbook of Nanotechnology, 3rd ed.;Bhushan, B., Ed.; Springer: New York, 2010; pp 1437–1524.
5. Rentschler, I. Planta 1971, 96, 119–135. doi:10.1007/BF003863626. Holloway, P. J. Pestic. Sci. 1970, 1, 156–163.
doi:10.1002/ps.27800104117. Neinhuis, C.; Barthlott, W. Ann. Bot. (Oxford, U. K.) 1997, 79, 667–677.
doi:10.1006/anbo.1997.04008. Mockenhaupt, B.; Ensikat, H. J.; Spaeth, M.; Barthlott, W. Langmuir
2008, 24, 13591–13597. doi:10.1021/la802351h9. Wagner, P.; Fürstner, R.; Barthlott, W.; Neinhuis, C. J. Exp. Bot. 2003,
54, 1295–1303. doi:10.1093/jxb/erg12710. Barthlott, W.; Neinhuis, C.; Jetter, R.; Bourauel, T.; Riederer, M.
Flora (Jena) 1996, 191, 169–174.11. Ensikat, H. J.; Boese, M.; Mader, W.; Barthlott, W.; Koch, K.
Chem. Phys. Lipids 2006, 144, 45–59.doi:10.1016/j.chemphyslip.2006.06.016
12. Ensikat, H. J.; Schulte, A. J.; Koch, K.; Barthlott, W. Langmuir 2009, 25,13077–13083. doi:10.1021/la9017536
13. Zhang, J.; Wang, J.; Zhao, Y.; Xu, L.; Gao, X.; Zheng, Y.; Jiang, L.Soft Matter 2008, 4, 2232–2237. doi:10.1039/b807857b
14. Cheng, Y. T.; Rodak, D. E.; Wong, C. A.; Hayden, C. A.Nanotechnology 2006, 17, 1359–1362.doi:10.1088/0957-4484/17/5/032
15. Zhang, J.; Sheng, X.; Jiang, L. Langmuir 2009, 25, 1371–1376.doi:10.1021/la8024233
16. Extrand, C. W.; Moon, S. I. Langmuir 2010, 26, 17090–17099.doi:10.1021/la102566c
17. Coward, J. L. J. Biol. Phys. 2010, 36, 405–425.doi:10.1007/s10867-010-9192-6
18. Mazliak, P. Chemistry of Plant Cuticles. In Progress in Phytochemistry,Vol. 1; Reinhold, L.; Liwschitz, Y., Eds.; Interscience Publishers: NewYork, 1968; pp 49–111.
19. Holloway, P. J. J. Sci. Food Agric. 1969, 20, 124–128.doi:10.1002/jsfa.2740200214
20. Leroy, F.; Müller-Plathe, F. Langmuir 2011, 27, 637–645.doi:10.1021/la104018k
21. Koch, K.; Ensikat, H. J. Micron 2008, 39, 759–772.doi:10.1016/j.micron.2007.11.010
22. Varanasi, K. K.; Deng, T.; Hsu, M.; Bhate, N. HierarchicalSuperhydrophobic Surfaces Resist Water Droplet Impact. InNanotechnology 2009: Biofuels, Renewable Energy, Coatings, Fluidicsand Compact Modeling;Nanoscience and Technology Institute, Cambridge, Ed.; TechnicalProceedings of the 2009 NSTI Nanotechnology Conference and Expo,Vol. 3; CRC Press: Boca Raton, 2009; pp 184–187.
23. Nearing, M. A.; Bradford, J. M.; Holtz, R. D. Soil Sci. Soc. Am. J. 1987,51, 1302–1306. doi:10.2136/sssaj1987.03615995005100050038x
24. Barthlott, W.; Schimmel, T.; Wiersch, S.; Koch, K.; Brede, M.;Barczewski, M.; Walheim, S.; Weis, A.; Kaltenmeier, A.; Leder, A.;Bohn, H. F. Adv. Mater. 2010, 22, 1–4. doi:10.1002/adma.201090075
25. Koch, K.; Barthlott, W. Philos. Trans. R. Soc., A 2009, 367, 1487–1509.doi:10.1098/rsta.2009.0022
26. Bush, J. W. M.; Hu, D. L.; Prakash, M. Adv. Insect Physiol. 2007, 34,117–192. doi:10.1016/S0065-2806(07)34003-4
27. Balmert, A.; Bohn, H. F.; Ditsche-Kuru, P.; Barthlott, W. J. Morphol., inpress. doi:10.1002/jmor.10921
28. Ensikat, H. J.; Ditsche-Kuru, P.; Barthlott, W. Scanning electronmicroscopy of plant surfaces: simple but sophisticated methods forpreparation and examination. In Microscopy: Science, Technology,Applications and Education; Méndez-Vilas, A.; Diaz, J., Eds.;FORMATEX Microscopy series No. 4, Vol. 1; Formatex ResearchCenter: Badajoz, Spain, 2010; pp 248–255.
29. Robinson, D. G.; Ehlers, U.; Herken, R.; Herrmann, B.; Mayer, F.;Schürmann, F. W. Präparationsmethodik in der Elektronenmikroskopie;Springer: Heidelberg, 1985; p 39.
30. Ensikat, H. J.; Neinhuis, C.; Barthlott, W.Int. J. Plant Sci. (Chicago, IL, U. S.) 2000, 161, 143–148.doi:10.1086/314234
31. Koch, K.; Dommisse, A.; Barthlott, W. Cryst. Growth Des. 2006, 6,2571–2578. doi:10.1021/cg060035w
License and TermsThis is an Open Access article under the terms of the
Creative Commons Attribution License
(http://creativecommons.org/licenses/by/2.0), which
permits unrestricted use, distribution, and reproduction in
any medium, provided the original work is properly cited.
The license is subject to the Beilstein Journal of
Nanotechnology terms and conditions:
(http://www.beilstein-journals.org/bjnano)
The definitive version of this article is the electronic one
which can be found at:
doi:10.3762/bjnano.2.19