an update on polygalacturonase- inhibiting protein (pgip), a … · 2020. 3. 17. · pgip gene...

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

[email protected]

Giulia De Lorenzo,

Dipartimento di Biologia e

Biotecnologie “Charles Darwin,”

Sapienza Università di Roma, Roma,

Italy

[email protected]

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

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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


http://www.ncbi.nlm.nih.gov/





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.






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


http://www.ncbi.nlm.nih.gov/nucleotide/





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


PVS-Fu 64

Fusarium sacchari

Fusarium fujikuroi

F. thapsinum


FC-10


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 sacchari

Fusarium fujikuroi

Fusarium verticillioides


ISPAVEmc 1189

Fusarium nygamai

Asparagus (Asparagus

officinalis L.)

White spear Mono-S

chromatography

Fusarium anthophilum Raiola et al., 2008

Fusarium circinatum

Fusarium subglutinans.


Fusarium sacchari

Fusarium fujikuroi

Fusarium verticillioides


ISPAVEmc 1189

Fusarium nygamai

Pepper (Capsicum

annuum L.)

Fruit Ion-exchange

chromatography

Colletotrichum

gleosporoides,

Shivashankar et al., 2010

Colletotrichum capsici,

Colletotrichum

lindemuthianum

Sclerotium rolfsi


(Continued)






TABLE 2 | Continued

Plant Tissue PGIP

preparation



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


Colletotrichum

lindemuthianum

Aspergillus niger

exopolygalacturonase

Ralstonia

solanacearum

(Continued)






TABLE 2 | Continued

Plant Tissue PGIP

preparation



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






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


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


Botrytis cinerea

Lima bean (Phaseolus

lunatus L.)

PlPGIP2 PVX/Nicotiana benthamiana Fusarium moniliforme§ Farina et al., 2009

Aspergillus niger


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


Botrytis aclada

Aspergillus niger

Botrytis cinerea

Colletotrichum acutatum

Fusarium graminearum

Lygus rugulipennis

Adelphocoris lineolatus

Orthops kalmi

Closterotomus norwegicus

(Continued)






TABLE 3 | Continued



GmPGIP3 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum PGb D’Ovidio et al., 2006; Frati



Botrytis aclada

Aspergillus niger

Botrytis cinerea



GmPGIP4 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum PGb D’Ovidio et al., 2006; Frati



Botrytis aclada

Aspergillus niger

Botrytis cinerea



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)






TABLE 3 | Continued



Raspberry (Rubus idaeus L.) RiPGIP Transgenic pea Stenocarpella maydis Richter et al., 2006


Wheat

(Triticum aestivum L.)

TaPGIP1

TaPGIP2

PVX/Nicotiana benthamiana Fusarium phyllophylu Janni et al., 2013

Stenocarpella maydis

Bipolaris sorokiniana


Rice (Oryza sativa L.) OsPGIP1 PVX/Nicotiana benthamiana Sclerotinia sclerotiorum Janni et al., 2006



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






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






(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






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.

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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.

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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

http://www.beilstein-journals.org/bjnano/about/openAccess.htm


http://dx.doi.org/10.3762%2Fbjnano.2.19

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.


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.


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


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°.


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


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-


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-


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.

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