Leman, A.

Pijnakker, J.

English

NARCIS 

Control of rose scale Aulacaspis Rosae (Bouché) (Hemiptera: Diaspididae) in greenhouse grown roses by releasing Rhizobius Lophantae (Blaisdell) (Coleoptera: Coccinellidae)

Name not available

Wageningen Yield 

AFPP – LES COCHENILLES : RAVAGEUR PRINCIPAL OU SECONDAIRE 
MONTPELLIER – 25 OCTOBRE 2011 
 
 
 
CONTROL OF ROSE SCALE AULACASPIS ROSAE (BOUCHÉ) (HEMIPTERA: 
DIASPIDIDAE) IN GREENHOUSE GROWN ROSES BY RELEASING RHIZOBIUS 
LOPHANTAE (BLAISDELL)  (COLEOPTERA: COCCINELLIDAE) 
  
J. PIJNAKKER & A. LEMAN 
 
Wageningen UR Glastuinbouw, Violierenweg 1, 2665 MV Bleiswijk, The Netherlands 
 
 
 
SUMMARY 
Problems with scale insects occur more frequently with the limited use of broad spectrum 
pesticides of the recent years. In Dutch greenhouses, scales are traditionally a problem in 
cymbidium, but since three years growers report increasingly the presence of Aulacaspis rosae 
(Bouché, 1933) in rose crops. Chemical control of scale insects is extremely difficult, because 
the pest spends most of its life hidden under its impenetrable shield. A short overview is given 
on the scale and on Integrated Pest Management methods: chemical control, spontaneously 
occurring parasitoids, first trials and releases of beneficials at growers. Biological control of the 
rose scale using the predatory beetle Rhizobius lophantae is now considered a promising 
alternative to chemical control. 
 
Key-words: scales, rose, greenhouse, Integrated Pest Management, natural enemies. 
 
 
RÉSUMÉ 
Depuis que les pesticides à large spectre d’action sont moins utilisés, les problèmes de 
cochenilles diaspines augmentent de façon préoccupante en cultures ornementales sous 
serre. Aux Pays-Bas, celles-ci sont des ravageurs courants en culture de Cymbidium, mais 
depuis trois ans la présence d’Aulacaspis rosae (Bouché, 1933) est de plus en plus rapportée 
en culture de fleurs coupées de roses. Le bouclier qui caractérise les cochenilles diaspines 
rend la protection chimique extrêmement difficile. Notre étude offre une synthèse rapide sur 
A. rosae et sur les méthodes de Protection Intégrée : protection chimique, ennemis naturels 
d’occurrence spontanée, premiers essais de protection intégrée et lâchés d’auxiliaires chez 
les producteurs. On considère actuellement les introductions de la coccinelle Rhizobius 
lophantae comme une alternative prometteuse à la protection chimique contre la cochenille 
de la rose. 
 
Mots-clés: cochenilles diaspines, rose, serre, Protection Intégrée, auxiliaires. 
 
 
INTRODUCTION 
Dutch rose growers in greenhouses ared faced with a new insect pest. A grower reported in 
2009 the presence of rose scale Aulacaspis rosae (Bouché) in his crop. The pest spread on 
the whole crop despite several pulverisations of all hot spots with broad spectrum 
insecticides and repeated drench applications of imidacloprid. In order to control the pest the 
grower had to stop the releases of beneficials and spray repeatedly the whole crop with non-
selective pesticides. Since then, growers are seeking to find solutions against this pest to 
limit its spread. 80% of the IPM rose growers have reported to have found the scale 
nowadays.   
At the request of the Dutch growers and with funding from the Dutch Product Board for 
Horticulture, Wageningen UR Greenhouse Horticulture conducts since 2010 experiments to 
improve the integrated pest control strategies against the rose scale. 
 
The rose scale  
The genus Aulacaspis belongs to the family Diaspididae. In the ancient world, this genus 
includes 29 species (Ben-Dov et al., 2010), of which only three are described in Europe: A. 
rosae (Bouche, 1833), A. tubercularis Newstead 1906 and A. yasumatsui Takagi 1977. 
Aulacapsis rosae is originally from Asia, and has been introduced to Europe, America, 
Australia and Africa (Ben-Dov et al., 2010). 
Aulacaspis rosae is known under the synonyms: Anamaspis rosae Kozarzhevskaya & 
Vlainic, 1981; Aulacaspis rosae Newstead, 1901; Diaspis rosae Froggatt, 1914; D. rosae 
Targioni Tozzetti, 1868; Aulacaspis rosae Cockerell, 1896; Aspidiotus rosae Bouché, 1833; 
Chermes rosae Boisduval, 1868. 
The rose scale is mainly found in the woody parts of rose plants. Besides roses, A. rosae is 
found on the following host plants: raspberries, logan berries, Rubus, black currant, 
Listroderes oblique, Agrimonia, Cycas, Dianthus, hydrangea, Laurus, Muehlenbeckia and 
Pyrus (http://nlbif.eti.uva.nl). Furthermore, its presence has been also reported on 
Anacardiaceae, Myricaceae and Saxifragaceae (Ben-Dov et al., 2010). 
The females of the rose scale are protected by a 1.5 to 2.5 mm shield. The shield is dirty 
white, flat and nearly round. The body of the female under the shield is long, orange to 
reddish brown in color and 1 mm long. The shield of the young males is oblong, white, flat, 
with a ribbed texture. Only the adult male is winged. His body is orange-red in color with a 
spine at the abdomen. Scale insects don’t secrete honeydew. 
The female lays between 50 and 150 eggs (Bazarov & Smelev, 1971) in a batch under the 
shield. The female dies shortly after oviposition. Hundreds of pale red to orange colored 
crawlers are leaving the shield. They settle on a suitable place and can not subsequently 
move anymore.  
The species has one (Sparrow, 1972) to four (Kosztarab & Kozar, 1988; Kozar, 1990) 
generations per year. The rose scale overwinters as eggs according to an American 
publication (Davidson & Peairs, 1966) or in all stages according to European authors 
(Bénassy, 1961, 1956; Kosztarab & Kozar, 1978). 
The crawlers are the only stage that contributes to active dispersal, but only on very small 
distance. The mortality is high in this phase. Passive dispersal occurs by transport on plants, 
humans, animals, and air movements. 
The rose scale causes direct damage by sucking sap from leaves, twigs and fruits, resulting 
in growth inhibition, discoloration of leaves and fruits, death of twigs and loss of production. 
At high density, the stalks and wood are covered with a white coating. Heavy infection can 
lead to the death of the whole plant (Kosztarab & Kozar, 1988). 
Young nymphs are easy to control, but the adult females are difficult to reach with 
insecticides; even more so if they are piled on each other. Broad-spectrum chemicals are 
especially effective. Pesticides have to be frequently sprayed with a lot of water with 7 - to 
14-day intervals. The addition of a wetting agent is recommended in order to optimize the 
coverage. Systemic compounds such as imidacloprid can affect older nymphal stages and 
adult females via the plant juices. Chemical insecticides effective against scale insects are 
acetamiprid, deltamethrin, dimethoate, imidacloprid, methiocarb, pyriproxyfen, spirodiclofen, 
spiromesifen, thiacloprid, and thiamethoxam. Some products of natural origin are also 
reported to be used against the pest: azadirachtin, pyrethrin and piperonyl butoxide, soaps, 
oils, algae extracts and products with physical mode of action.  
In roses, the scale insects are primarily found on the old wood, in the heart of the plant or 
lower parts of the plants (bent shoots). But by high infestation levels crawlers reach the 
flower buds. The effects of plant protection products, even if they work systemically, is often 
disappointing due to the limited coverage. Growers need to spray carefully for an optimal 
coverage to get between the bent shoots.  
To avoid the establishment of the crawlers some authors recommend the application of 
mineral oil. Some Dutch growers cover the foot of the plant with a coating balsam after they 
have treated them with an insecticide.  
Beetles are the most studied predators of scales, and more than 20 species of parasitoids 
have been identified in association with the rose scale. Two of them: Arrhenophagus 
chionaspidis Aurivillius (Hymenoptera, Encyrtidae) and Adelencyrtus aulacaspidis (Brèthes) 
(Hymenoptera, Encyrtidae) occur spontaneously in some Dutch greenhouses. Our research is 
focussing on testing beneficials and the combination of beneficials and selective chemicals.   
 
MATERIAL AND METHODS 
To test the efficacy of four beneficials against the rose scale an experiment was set up in a 
randomized complete block design with five treatments and three replications in a 
greenhouse at the research station. The trial was conducted from February to May 2011.  
Fifteen experimental cages of dimensions of 1 x 1.5 x 1 m with a metal frame were used. 
They were covered with fine mesh screen with a 0.22 x 0.31 mm opening size. Each cage 
contained three rose plants cv. Wham provided by Schreurs (Aalsmeer, NL) and had two 
zippers through which plants could be handled or examined.  
The cages were distributed on eight tables in a greenhouse of 144 m² with 70% RH and a 
temperature fluctuating around 22°C.  
A. rosae colonies were collected at a grower from Stompwijk (NL). The plants were infested 
with scales on February4th.  
Inundative introductions of Amblyseius swirskii Athias-Henriot (Acari, Phytoseiidae), 
Eretmocerus eremicus Rose & Zolnerowich (Hymenoptera, Aphelinidae) and Phytoseiulus 
persimilis Athias-Henriot (Acari, Phytoseiidae), were regularly carried out to prevent any 
establishment of thrips, whiteflies or spider mites. 
Treatments were blocked according to a precount of the scale populations on the plants in 
week 14 on April 6th (Block 1: average 210 scales/cage, Block 2: 1300, Block 3: 12300). The 
presence of A. chionaspidis was detected prior to the release of the natural enemies. The first 
beneficials were released on the same day. The natural enemies used in this study were 
supplied by Entocare CV., except the parasitoid A. chionaspidis which was reared at 
Wageningen UR Greenhouse horticulture. The introduction rates were chosen in 
collaboration with the supplier of beneficials (Table I).  
 
Table I: Introduction of beneficials 
            (Introduction d’ auxilliaires) 
Predator 
Supplier Number of 
introductions 
Quantity 
Untreated - - - 
Karnyothrips melaleucus  
Entocare CV., 
Wageningen, NL 
4 
50 adults /cage/week 
Arrhenophagus chionaspidis 
Wageningen UR 
Greenhouse Horticulture 
Bleiswijk, NL 
4 
30 adults/cage/week 
Rhizobius lophantae 
Entocare CV., 
Wageningen, NL 
4 
15 adults/cage/2 weeks 
Encarsia citrina  
Entocare Cv., 
Wageningen, NL 
4 
30 adults /cage/week 
 
Assessments 
The counts of scales per cage were recorded on the day of the first introduction and four, 
seven and ten weeks after the first introduction of biological agents. The number of predators 
per cage was assessed with a magnifier four, seven and ten weeks after the first introduction 
of biological agents. Parasitism rate was studied at three data on 100 nymphs of the second 
stage.  
 
In order to evaluate the presence of Encarsia citrina Craw (Hymenoptera, Aphelinidae), 100 
pupae from each cage were transferred at three data in ventilated plastic containers (8.5 cm 
in diameter and 6 cm high, with a ventilation hole of 3 cm in diameter made in the lid and 
covered with thrips-proof nylon gauze). A 4 x 4 cm section of a yellow sticky trap (Horiver, 
Koppert, NL) was placed in the container to trap emergent adults. The containers were kept 
at 20ºC, 80% r.h., and L16:D8 photoperiod during one week. One yellow sticky trap was 
hung during one week in the cages at the end of the experiment. 
 
Data analysis  
An ANOVA using the Generalized Linear Model procedure with binomial distribution was 
performed for the test on parasitism. A generalized linear model with a Poisson distribution 
was used for the analyses and the number of scales in the pre-counting was taken into 
account. For pairwise comparisons of treatment means a likelihood ratio test was used. 
 
RESULTS AND DISCUSSION 
During the experiment, the reproduction of the beetle was poor: only one or two larvae of R. 
lophantae were found per assessment in the three cages. Karnyothrips melaleucus (Bagnall) 
(Thysanoptera, Phlaeothripidae) didn’t establish at all. E. citrina was found sporadically at the 
end of the experiment (0.7, 2 and 4% of the emerged parasitoids respectively in week 18, 22, 
24). Only A. chionaspidis was present in all the cages except in cages were R. lophantae 
was released. The beetle predated on all parasitized scales.  
 
On May 3rd, the beetle had already predated 70 % of the scales (Figure 1). In all other 
treatments, the amount of scales increased despite the introductions of beneficials and the 
natural presence of A. chionaspidis. The pest continued to spawn until the end of the 
experiment, except in the cages with R. lophantae where they were almost eradicated.   
 
Parasitism rates varied between 17 and 49 % (Figure 2). A. chionaspidis was the predominant 
species. No significant difference was observed between the treatments. The release of 
parasitoids had no additional value.  
Figure 1: Scale density per treatment (Km: Karnyothrips melaleucus, Ar: Arrhenophagus 
chionaspidis, Rl: Rhizobius lophantae and Ec: Encarsia citrina) 
                [Densité de cochenilles par modalité (Km: Karnyothrips melaleucus, Ar: 
Arrhenophagus chionaspidis, Rl: Rhizobius lophantae et Ec: Encarsia citrina)] 
 
 
 
Figure 2: Parasitism rates (Km: Karnyothrips melaleucus, Ar: Arrhenophagus chionaspidis, 
Rl: Rhizobius lophantae and Ec: Encarsia citrina) 
                [Taux de parasitisme (Km: Karnyothrips melaleucus, Ar: Arrhenophagus 
chionaspidis, Rl: Rhizobius lophantae et Ec: Encarsia citrina)] 
 
 
Implementation in commercial greenhouses  
These results are confirmed with field experience of growers. The scale spreads in the whole 
crop even with high parasitism rates (60-80%) of A. chionaspidis. About 100 R. lophantae were 
introduced by a grower in May 2010 on few hot spots. The beetle was able to maintain itself in 
the crop during the whole year with rates of 1/20 infested plants during the winter and densities 
increased to 3 to 9/infested plant from March on. The rose scale dispersed widely in the crop in 
November, but in April the grower couldn’t find any new hot spots. The pest was under control in 
June. The monitoring of the pest and its natural enemies continues.  
Another grower introduces the beetle at a rate of 20 beetles per new hotspot of about 50 scales 
and is keeping the pest under control until now. The results so far with this predator look very 
promising. 
 
OUTLOOKS 
Most of Dutch growers chose in 2010 to frequently spray the hot spots with neonicotinoids, 
but they often ended the year with applications of pesticides in their whole crop.  
Growers will have to improve their scouting methods to detect the hot spots in an early stage 
or to apply IPM methods to control the rose scale. More growers are now introducing R.  
lophantae in combination with applications of selective chemicals to restrain the proliferation 
of the pest.  
 
ACKNOWLEDGEMENTS 
The authors gratefully acknowledge the participation of several greenhouse rose growers 
and we thank all the Dutch growers who provided financial support. Our thanks are due to 
Andrew Polaszek and John Noyes at the National History Museum of London for confirming 
the identification of the parasitoids. We also thank Irene Vanninen for her comments.  
 
REFERENCES  
Bazarov B.B. & Smelev G.P., 1971 - Armoured scales of Tadzhikistan and the adjoining regions 
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limitation des pullulations de cochenille-diaspines. Annales des Epiphyties, 12, 1-157. 
Bénassy C., 1956 - No title. Bull. Soc. Ent. Fr., 61, 5-6, 103-106. 
Ben-Dov Y., Miller D.R. & Gibson G.A.P., 2010 - ScaleNet: a database of the scale insects of 
the world. Available from http://www.sel.barc.usda.gov/scalenet/scalenet.htm. 
Davidson R.H. & Peairs L.M., 1966 - Insect pests of farm, garden and orchard. Fifth edition. 
John Wiley and Sons, New York, USA, 675 pp. 
Kosztarab M. & Kozár F., 1978 - Pajzstetvek - Coccoidea. (In Hungarian, with English 
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Control of rose scale Aulacaspis Rosae (Bouché) (Hemiptera: Diaspididae) in greenhouse grown roses by releasing Rhizobius Lophantae (Blaisdell) (Coleoptera: Coccinellidae)

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