Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide


Armillaria novae-zelandiae



Armillaria novae-zelandiae


  • Last modified
  • 14 February 2020
  • Datasheet Type(s)
  • Invasive Species
  • Preferred Scientific Name
  • Armillaria novae-zelandiae
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Fungi
  •     Phylum: Basidiomycota
  •       Subphylum: Agaricomycotina
  •         Class: Agaricomycetes
  • Summary of Invasiveness
  • Armillaria novae-zelandiae is a white rot wood decay fungus and root disease pathogen that occurs in a number of countries in the Southern Hemisphere and in parts of tropical and subtropical Asia. It is not kno...

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Armillaria novae-zelandiae; habit. Fox Glacier, West Coast, New Zealand. April 2010.
CaptionArmillaria novae-zelandiae; habit. Fox Glacier, West Coast, New Zealand. April 2010.
CopyrightPublic Domain - Released by Bernard Spragg/via flickr - CC0
Armillaria novae-zelandiae; habit. Fox Glacier, West Coast, New Zealand. April 2010.
HabitArmillaria novae-zelandiae; habit. Fox Glacier, West Coast, New Zealand. April 2010.Public Domain - Released by Bernard Spragg/via flickr - CC0


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Preferred Scientific Name

  • Armillaria novae-zelandiae (G. Stevenson) Herink

Other Scientific Names

  • Agaricus melleus sensu Cooke
  • Armillaria mellea sensu Massee
  • Armillaria novae-zelandiae (G. Stevenson) Boesewinkel
  • Armillariella mellea sensu G. Stevenson
  • Armillariella novae-zelandiae G. Stevenson
  • Armillariella sparrei var. elaeodes Singer

Local Common Names

  • New Zealand: harore

Summary of Invasiveness

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Armillaria novae-zelandiae is a white rot wood decay fungus and root disease pathogen that occurs in a number of countries in the Southern Hemisphere and in parts of tropical and subtropical Asia. It is not known to have been introduced to these regions, where it is presumed to be indigenous. Its designation as “invasive” is based on its propensity to establish colonies and disease centres in disease-free areas by dispersal of basidiospores from “toadstool” fruit bodies that appear on wood during the winter months. As a wood decomposer fungus A. novae-zelandiae contributes beneficially to carbon and nutrient recycling. Like many other Armillaria species it is recognized by characteristic white mycelial fans or ribbons produced beneath host bark and by its bootlace-like rhizomorphs by which it spreads vegetatively from colonized buried woody material or stump root systems to infect living host plants.

Armillaria novae-zelandiae was the cause of substantial disease losses in plantations of Pinus radiata and orchards of kiwifruit vines (Actinidia deliciosa) in New Zealand from the 1970s to the 1990s. Its importance has since declined with changes in patterns of crop management, although it remains widely distributed. Much research into its control was undertaken during this period. In eastern states in Australia, Anovae-zelandiae is a minor cause of root disease in natural and planted forests, where it is of lesser importance than Armillaria luteobubalina. Its impact in other regions is unknown, but it has not been associated with reports of significant disease. Risk of unintended international spread appears to be low to negligible but should not be discounted. If intercepted, isolates of Anovae-zelandiae may be identified by laboratory culture testing or more rapidly and precisely by molecular sequencing procedures. Armillaria novae-zelandiae is listed in the EPPO Global Database and features in the United States Department of Agriculture Agricultural Research Service fungal databases. It is considered a risk organism in Hawaiʻi.

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Fungi
  •         Phylum: Basidiomycota
  •             Subphylum: Agaricomycotina
  •                 Class: Agaricomycetes
  •                     Subclass: Agaricomycetidae
  •                         Order: Agaricales
  •                             Family: Marasmiaceae
  •                                 Genus: Armillaria
  •                                     Species: Armillaria novae-zelandiae

Notes on Taxonomy and Nomenclature

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Armillaria novae-zelandiae is a well-defined species that demonstrates monophyletic, within-species molecular variation. Isolates from different geographic regions tend to group separately within the same clade, based on phylogenetic analysis of DNA sequences from the internal transcribed regions (ITS1, 5.8S and ITS4) and ribosomal large subunit (RSU) (Coetzee et al., 2003a, Pildain et al., 2009; 2010, Ota et al., 2011). This separation is not considered sufficient to divide Anovae-zelandiae into separate within-species taxa or to erect new species. The uniqueness of Anovae-zelandiae as a species is supported by phylogenetic analysis using the partial elongation factor 1-alpha gene region (Maphosa et al., 2006) and by interfertility studies using haploid isolates (Kile and Watling, 1988).

Pildain et al. (2010) synonymized Asparrei var. elaeodes with Anovae-zelandiae. In New Zealand, Armillariella mellea sensu G. Stevenson, Agaricus melleus sensu Cooke and Armillaria mellea sensu Massee were used for both A. novae-zelandiae and Armillaria limonea (G. Stevenson) Boesewinkel before it was accepted that Armillaria mellea (Vahl) Kumm. is not naturally present in the Southern Hemisphere. The Maori word harore is a term used especially for the common New Zealand Armillaria species fruiting in winter, but also loosely for other edible agarics or fungi in general.

The holotype of A. novae-zelandiae, collected by G. Stevenson in New Zealand in 1949, is in the Fungarium at the Royal Botanic Gardens, Kew (Kew Mycology Collection, 2020).


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Descriptions of Armillaria novae-zelandiae are provided by Stevenson (1964), Singer (1969), Kile and Watling (1983) and Gadgil (2005). See also van der Pas et al. (2008) and Cooper (2016).

As described by Gadgil (2005): “Basidiomata pileate, centrally stipitate. Pileus 30–80 mm in diameter, olive buff to olive brown, sprinkled with very small dull brown scales at the centre, convex at first, becoming plane to shallow concave; flesh creamy white. Gills decurrent to almost sinuate, moderately crowded, creamy white becoming dull fawn. Stipe tapering towards the base, fawn above evanescent annulus, brown to dark purplish below, smooth or striate, solid, 40–70 mm long. Basidiospores ellipsoid to elongate-ellipsoid, 7–9 × 5–6 m, non-amyloid; spore print white”.

The cap is initially viscid (tacky). Rhizomorphs are common, dichotomous in form (Benjamin, 1983; Morrison, 1989).


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Armillaria novae-zelandiae is found in Australasia (including Papua New Guinea), southern South America and southeastern Asia (Indonesia, Malaysia and the Amami Archipelago in southwestern Japan). It shows significant regional molecular variation between isolates from Australasia, Asia (Malaysia, Sumatra, Japan) and South America (Coetzee et al., 2003a; Pildain et al., 2009; 2010; Ota et al., 2011). Armillaria, as A. mellea (with “strong black rhizomorphs”; Parham 1953) and as Clitocybe tabescens (in plantations of Pinus caribaea var. hondurensis; Singh, 1978), occurs in Fiji, but its precise identity is unknown and the presence of A. novae-zelandiae remains to be confirmed.

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Last updated: 14 Feb 2020
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes


-SumatraPresentNative2003Coetzee et al. (2003)Lake Toba district
JapanPresentCABI (2020)Present, based on regional distribution
-Ryukyu IslandsPresentNative2011Ota et al. (2011)Island of Amami-Oshima
MalaysiaPresentNative2003Coetzee et al. (2003)


AustraliaPresentCABI (2020)Present, based on regional distribution
-New South WalesPresentNative1988Kile and Watling (1988)One of five Armillaria species indigenous to Australia
-QueenslandPresentNative1988Kile and Watling (1988)One of five Armillaria species indigenous to Australia. Occurs in southern Queensland as far north as the Bunya Mountains
-TasmaniaPresentNative1983Kile and Watling (1988)One of five Armillaria species indigenous to Australia
-VictoriaPresentNative1983Kile and Watling (1988)One of five Armillaria species indigenous to Australia
FijiAbsent, Unconfirmed presence record(s)Baumgartner et al. (2011)Requires confirmation from an original literature source
New ZealandPresent, WidespreadNative1964InvasiveStevenson (1964)Present throughout New Zealand, including the Chatham Islands. One of four Armillaria species indigenous to New Zealand
Papua New GuineaPresentNative1992Guillaumin et al. (1992)Mount Giluwe and Mount Wilhelm upper altitude regions. Armillaria spp. (but not specifically A. novae-zelandiae) are reported earlier in PNG than 1992

South America

ArgentinaPresentNative1969Singer (1969)Patagonian Andes (Neuquén) and Buenos Aires Province
ChilePresentNative1969Singer (1969)Llanquihue and Isla Grande de Chiloé

History of Introduction and Spread

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There is no evidence that A. novae-zelandiae has spread internationally due to human activity. It is apparently indigenous in all places where it has been found, due to ancient basidiospore dispersal probably after the breakup of Gondwana (Coetzee et al., 2011; 2018; Klopfenstein et al., 2017; cf. distribution of certain other agaric species, Horak, 1983).

Risk of Introduction

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There are several documented accounts of the probable spread of Northern Hemisphere Armillaria species by human means. In Western Cape province in South Africa. Armillaria mellea was found in woody ornamental trees and shrubs in public gardens in Cape Town (Coetzee et al., 2001a) and subsequently A. mellea and A. gallica were both identified infecting Protea and Leucadendron plants in another public garden in the same Province (Coetzee et al., 2003b). Both introductions were considered to date back to early Dutch settlement by means of infested soil with potted plants. Later evidence has suggested that human movement of plant material from Japan, or from China via Japan, is responsible for the introduction of homothallic (self-fertile) populations of Amellea to western and eastern Africa (Baumgartner et al., 2012; Guo et al., 2016). From the extent of their distribution, it appears that these introductions were also not recent. Both Amellea and A. gallica are reported as introduced into Hawaiʻi, where they are threatening native vegetation, the former possibly on nursery fruit stock in the early 1900s and the latter more recently (Burgan and Nelson, 1972; Gardner, 2003; Kim et al., 2010; cf. Raabe et al., 2009).

Despite these reports, the risk of A. novae-zelandiae spreading internationally appears low to negligible, however it cannot be ignored. Armillaria novae-zelandiae is largely confined to roots and root collars of infected trees and shrubs. In young plants mycelial fans may extend above soil level only if host resistance declines prior to or after death, or because of physiological stress resulting from some additional external physical, chemical or biological cause. In older trees A. novae-zelandiae may form a butt rot or basal canker immediately above ground level. However, though possible, carriage on eucalypt logs from eastern Australia is considered of low risk (Kliejunas et al., 2003). The export of old growth native timber from New Zealand is restricted (Forestry New Zealand, 2020). In a proportion of mature Pradiata trees (up to ca. 25 years old) non-lethal by Anovae-zelandiae (see below) is present only at and below soil level, indicating a zero to negligible risk of its being present in harvested logs cut at stump height (butt rot does not develop prior to harvest in these trees). Armillaria novae-zelandiae requires a supporting wood- or organic-based substrate, so its presence in nurseries is rare. Any infected plants would normally show readily detectable disease symptoms. It is possible that soil admixtures such as peat or other organic material might support A. novae-zelandiae saprotrophically if the pathogen was somehow introduced, but the risk of movement in nursery stock would seem low.

Discussion on the risk of introducing Armillaria species to North America on eucalypt logs and chips from Australia is included in Kliejunas et al. (2003) and from South America in Kliejunas et al. (2001). Similar information and discussion regarding the movement of Pinus radiata and Douglas fir logs from New Zealand to North America can be found in White et al. (1992). Armillaria novae-zelandiae is listed in the EPPO Global Database (EPPO, 2019) and features in the United States Department of Agriculture Agricultural Research Service fungal databases (USDA-ARS, 2019). Armillaria novae-zelandiae is considered to be a risk organism in Hawaiʻi (DeNitto et al., 2015).


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Armillaria novae-zelandiae occurs naturally in woody vegetation of various kinds. Available information on the occurrence of A. novae-zelandiae in South America, Asia or Oceania is limited. In South America it was found on living and dead trunks of dicotyledenous trees such as Nothofagus and Salix species (Singer, 1969). In subtropical southern Japan it was reported in evergreen forest dominated by Castanopsis sieboldii, Quercus miyagi and Distylium racemosum (along with planted Pinus luchuensis) (Ota et al., 2011). In Australia it occurs in all four States (including Tasmania) along the eastern side of the continent on standing dead trees, stumps or fallen trees or shrubs of species such as Eucalyptus spp., Lophozonia (Nothofagus) cunninghamii, Atherosperma moschatum, Acacia melanoxylon, Pomaderris sp., Olearia sp. and Phebalium sp. (Kile and Watling, 1983; 1988). It is apparently more common in montane native forests in this region (Kile and Watling, 1988).

In New Zealand, A. novae-zelandiae occurs naturally (along with A. limonea) as a very common saprotrophic fungus causing decay in coarse woody debris and as a butt rot species in living trees in native forests, both podocarp-hardwood and southern beech (Nothofagus) (Colenso, 1890; Birch, 1937a; Gilmour 1954; 1966; Hood and Sandberg, 1987; Hood et al., 2004c; 2019). Rhizomorphs are prolific in the soil beneath podocarp-hardwood forests, growing epiphytically across healthy roots without infecting them (roots of older trees can be found with Armillaria-caused decay, but it is not clear whether the fungus is the cause of death or if the roots were killed or weakened from some other cause). Armillaria novae-zelandiae is not generally seen as a pathogen of native trees (Rawlings, 1953) but Armillaria-caused mortality has occurred e.g., in densely stocked Nothofagusmenziesii saplings regenerating in gaps after storm damage (Birch, 1937b) and in the same host planted as an ornamental specimen tree in urban settings (Hood, 1989). Nevertheless, Armillaria species provide an indispensable service as important members of the indigenous wood decay community. Along with other decomposer fungi, A. novae-zelandiae contributes significantly to carbon and nutrient recycling in the native forest ecosystem (Hood et al., 2019).

Habitat List

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Terrestrial – ManagedManaged forests, plantations and orchards Secondary/tolerated habitat Harmful (pest or invasive)
Urban / peri-urban areas Present, no further details Harmful (pest or invasive)
Terrestrial ‑ Natural / Semi-naturalNatural forests Principal habitat Natural
Scrub / shrublands Present, no further details Natural

Hosts/Species Affected

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Armillaria novae-zelandiae occurs as a parasite on a wide range of host species, both angiosperms and gymnosperms. An online global list of hosts is provided by the Agricultural Research Service, United States Department of Agriculture (USDA-ARS, 2019). Host lists for New Zealand are given by Gilmour (1966); Dingley (1969); Laundon (1973); Shaw et al. (1976); Pennycook (1989); McKenzie et al. (1999; 2000; 2002); Gadgil (2005) and van der Pas et al. (2008) (A. novae-zelandiae and A. limonea are not always distinguished in these lists).

In New Zealand Armillaria species (including A. novae-zelandiae) are or have been important in fruit orchards (Atkinson, 1971), kiwifruit orchards (Actinidiadeliciosa), riparian willows (Salix species) and in forest plantations, particularly of Pinus radiata. Pathogenicity by A. novae-zelandiae to this host has been demonstrated by inoculation experiments with seedlings (Shaw et al., 1981; Benjamin, 1983; Benjamin and Newhook, 1984b; Hood and Sandberg, 1993b). Some of the minor scattered damage in pine and eucalypt plantations along the eastern States of Australia is due to A. novae-zelandiae (e.g., in a first rotation Pradiata plantation in southern Queensland; Kile and Watling, 1988). In Papua New Guinea there are records of Armillariamellea” in crops of Camellia sinensis, Coffea spp., Musa spp., and Theobroma cacao L., but it is not clear how much of this, if any, is due to A. novae-zelandiae or to other Armillaria species (Shaw, 1984). Similarly, minor damage due to Armillaria is reported in pines (including P. radiata) and other crops in southern South America, but the degree to which A. novae-zelandiae is implicated is unknown. Brief reviews of Armillaria in planted hosts in South America, Australasia and Papua New Guinea can be found in Hood et al. (1991), generally without indication of the contribution made by A. novae-zelandiae.

Host Plants and Other Plants Affected

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Plant nameFamilyContext
Acacia mangium (brown salwood)FabaceaeUnknown
Acacia melanoxylon (Australian blackwood)FabaceaeUnknown
Actinidia deliciosa (kiwifruit)ActinidiaceaeMain
Agathis australis (kauri)AraucariaceaeUnknown
Aristotelia serrataGelechiidaeUnknown
Atherosperma moschatumMonimiaceaeUnknown
Austroderia fulvidaUnknown
Bambusa oldhamiiPoaceaeUnknown
Beilschmiedia tawaLauraceaeUnknown
Buddleia (Butterflybush)LoganiaceaeUnknown
Buddleja davidii (butterfly bush)LoganiaceaeUnknown
Casuarina cunninghamiana (Australian beefwood)CasuarinaceaeUnknown
Cedrus atlantica (Atlas cedar)PinaceaeUnknown
Chamaecyparis lawsoniana (Port Orford cedar)CupressaceaeUnknown
Citrus sinensis (navel orange)RutaceaeUnknown
Cryptomeria japonica (Japanese cedar)TaxodiaceaeUnknown
Cupressus macrocarpa (Monterey cypress)CupressaceaeUnknown
Dacrycarpus dacrydioides (white pine)PodocarpaceaeUnknown
Dacrydium cupressinumPodocarpaceaeUnknown
Discaria toumatouRhamnaceaeUnknown
Eucalyptus delegatensis (alpine ash)MyrtaceaeUnknown
Eucalyptus grandis (saligna gum)MyrtaceaeUnknown
Eucalyptus regnans (mountain ash)MyrtaceaeUnknown
Eucalyptus spp.MyrtaceaeOther
Euonymus japonicus (Japanese spindle tree)CelastraceaeUnknown
Grevillea robusta (silky oak)ProteaceaeUnknown
Griselinia littoralisCornaceaeUnknown
Hedycarya arboreaMonimiaceaeUnknown
Jacaranda mimosifolia (jacaranda)BignoniaceaeUnknown
Knightia excelsaProteaceaeUnknown
Larix decidua (common larch)PinaceaeUnknown
Larix kaempferi (Japanese larch)PinaceaeUnknown
Laurelia novae-zelandiaeMonimiaceaeUnknown
Leycesteria formosaCaprifoliaceaeUnknown
Liriodendron tulipifera (tuliptree)MagnoliaceaeUnknown
Litsea calicarisLauraceaeUnknown
Lycium ferocissimum (African boxthorn)SolanaceaeUnknown
Malus domestica (apple)RosaceaeUnknown
Manoao colensoiPodocarpaceaeUnknown
Meryta sinclairiiUnknown
Metasequoia glyptostroboides (water fir)TaxodiaceaeUnknown
Metrosideros kermadecensisMyrtaceaeUnknown
Nothofagus alpina (rauli beech)NothofagaceaeUnknown
Nothofagus dombeyi (coigue)NothofagaceaeUnknown
Nothofagus fusca (red beech)NothofagaceaeUnknown
Nothofagus menziesiiNothofagaceaeUnknown
Nothofagus obliqua (roble)NothofagaceaeUnknown
Nothofagus pumilioNothofagaceaeUnknown
Nothofagus solandriNothofagaceaeUnknown
Nothofagus solandri var. cliffortioidesNothofagaceaeUnknown
Nothofagus truncataNothofagaceaeUnknown
Paraserianthes lophantha (brush wattle)FabaceaeUnknown
Persea americana (avocado)LauraceaeUnknown
Phyllocladus alpinusPodocarpaceaeUnknown
Pinus caribaea (Caribbean pine)PinaceaeUnknown
Pinus contorta (lodgepole pine)PinaceaeUnknown
Pinus elliottii (slash pine)PinaceaeUnknown
Pinus muricata (bishop pine)PinaceaeUnknown
Pinus nigra ssp. laricioPinaceaeUnknown
Pinus patula (Mexican weeping pine)PinaceaeUnknown
Pinus pinaster (maritime pine)PinaceaeUnknown
Pinus ponderosa (ponderosa pine)PinaceaeUnknown
Pinus radiata (radiata pine)PinaceaeMain
Pinus strobus (eastern white pine)PinaceaeUnknown
Pinus taeda (loblolly pine)PinaceaeUnknown
Pittosporum crassifoliumPittosporaceaeUnknown
Podocarpus totara (totara)PodocarpaceaeUnknown
Populus sp. (poplar)SalicaceaeUnknown
Prumnopitys taxifoliaPodocarpaceaeUnknown
Prunus armeniaca (apricot)RosaceaeUnknown
Prunus persica (peach)RosaceaeUnknown
Pseudotsuga menziesii (Douglas-fir)PinaceaeUnknown
Pyracantha crenulataRosaceaeUnknown
Pyrus communis (European pear)RosaceaeUnknown
Rhododendron (Azalea)EricaceaeUnknown
Ribes nigrum (blackcurrant)GrossulariaceaeUnknown
Ribes uva-crispa (gooseberry)GrossulariaceaeUnknown
Rubus idaeus (raspberry)RosaceaeUnknown
Salix (willows)SalicaceaeUnknown
Salix alba var. vitellinaUnknown
Salix fragilis (crack willow)SalicaceaeUnknown
Salix matsudana (Peking willow)SalicaceaeUnknown
Schefflera digitataAraliaceaeUnknown
Tecoma capensis (Cape honeysuckle)BignoniaceaeUnknown
Thuja plicata (western redcedar)CupressaceaeUnknown
Tsuga heterophylla (western hemlock)PinaceaeUnknown
Vitis vinifera (grapevine)VitaceaeUnknown
Weinmannia racemosa (maori)CunoniaceaeUnknown

Growth Stages

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As with other Armillaria species, plants infected by A. novae-zelandiae suffer injury and death of tissues in the roots and root collar region, accompanied by resinosis or gummosis (depending on the host). Characteristic signs are black, shoelace-like rhizomorphs and white mycelial sheeting or ribbons beneath the bark. Younger plants wilt or show crown discoloration and retain dead foliage for a time if they are killed rapidly. Older trees may decline more slowly or, in the case of Pinus radiata, retain green foliage, to all intents and purposes appearing healthy, although infected at the root collar. Old, mature trees may develop butt rot.

List of Symptoms/Signs

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SignLife StagesType
Leaves / abnormal colours
Leaves / wilting
Leaves / yellowed or dead
Roots / 'dirty' roots
Roots / fungal growth on surface
Roots / reduced root system
Roots / rot of wood
Stems / gummosis or resinosis
Stems / internal discoloration
Stems / mycelium present
Stems / rot
Whole plant / discoloration
Whole plant / dwarfing
Whole plant / plant dead; dieback
Whole plant / uprooted or toppled
Whole plant / wilt

Biology and Ecology

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Like many other Armillaria species, A. novae-zelandiae is assumed to have a diploid vegetative mycelium, undergoing meiosis to form haploid basidiospores (the basidium lacks a basal clamp connection; Kile and Watling, 1983).

Inherently, A. novae-zelandiae appears to show an intermediate level of virulence when compared with other saprotrophic and pathogenic species of Armillaria. In Australia it has less impact than A. luteobubalina. In New Zealand, A. novae-zelandiae has not shown secondary spread between adjacent root systems of young infected Pinus radiata trees as occurs, for instance, with the more virulent Armillariaostoyae in Pinus pinaster stands in the Landes de Gascogne forest region in southwest France (Roth et al., 1979; van der Pas, 1981b; Lung-Escarmant and Guyon, 2004). Pathogenicity is influenced environmentally as well as genetically: Armillaria, probably including A. novae-zelandiae, is observed to acquire greater inoculum potential on hardwood stumps of indigenous or exotic host species than on pine stumps, causing greater mortality in young, newly established P. radiata plantations. Inoculation studies have demonstrated that different A. novae-zelandiae isolates vary innately in their level of virulence to Pinus radiata (Shaw et al., 1981; Benjamin, 1983; Hood and Sandberg, 1993b; Hood et al., 2009).

Reproductive Biology

Fruiting occurs over several weeks in the Southern Hemisphere’s autumn and winter, between March and May (South America, Australia; Singer, 1969; Kile and Watling, 1983) or April to July (New Zealand; Hood et al., 2004b; Hood and Gardner, 2005). In New Zealand small and often very large clusters of fruitbodies appear along with those of Armillaria limonea in natural forests and on woody debris in indigenous forests, producing copious quantities of basidiospores. Fruiting is observed less commonly on infected introduced host trees and shrubs. There is no asexual stage.

Physiology and Phenology

Plantation P. radiata trees older than ca. 10 years (near mid-rotation) infected by Anovae-zelandiae are not killed but interact dynamically with the pathogen while remaining green and apparently healthy. Over time, infection at the root collar decreases in some trees while increasing in others (MacKenzie, 1987; Hood et al., 2002c). A. novae-zelandiae induces abnormal production of bark in the root collar region in which it survives and from which it penetrates and causes small lesions in the cambium during the cooler seasons when trees are less active (Gilmour, 1954; van der Kamp and Hood, 2002). These lesions heal during the warmer, more active growth periods but there is possibly an energy cost to the host associated with keeping the pathogen at bay.

Armillaria limonea, and presumably also A. novae-zelandiae, demonstrate bioluminescence and in New Zealand wood decayed by Armillaria may be seen faintly glowing in native forests on dark nights. Cultures are also bioluminescent, especially when young and actively growing.

Population Size and Structure

Populations of A. novae-zelandiae in New Zealand are composed of comparatively high colony densities per unit ground area. In a study in natural podocarp hardwood forest in New Zealand, cultural pairing of isolates demonstrated a density of colonies (vegetative compatibility groups) of A. novae-zelandiae estimated at between 19 and 93/ha (Hood and Sandberg, 1987; Hood, 1989). Similar studies in young P. radiata plantations that had replaced native forest yielded equivalent values of 32 colonies/ha (Benjamin, 1983; Benjamin and Newhook, 1984a) and between 46 and 108 colonies/ha (Hood and Sandberg, 1993b).

Rhizomorphs are produced prolifically (Farr et al., 1919). In New Zealand Hood and Sandberg (1987) recorded an average aggregate rhizomorph length (comprising both A. novae-zelandiae and A. limonea) ranging from 2 to 9 m/m2 of soil surface down to a depth of 22 cm beneath a podocarp-hardwood forest logged of the podocarp element three decades earlier.


In New Zealand, an Armillaria species that is probably A. novae-zelandiae (incorrectly described in the paper as A. mellea, which is not found in the Southern Hemisphere), forms a mycorrhizal relationship with the native orchid, Gastrodia cunninghamii in beech forests (Campbell, 1962; cf. Cone, 1959). Armillaria novae-zelandiae appears to behave similarly in Pinus radiata plantations with G. cunninghamii, Gastrodia aff. sesamoides, or both, in effect apparently facilitating indirect parasitism of the pines by the orchid species (Hood et al., 2004a; Hood, 2005). In New Zealand’s North Island podocarp-hardwood forest, Anovae-zelandiae has been found colonizing dead plants of the root parasite Dactylanthus taylori (Balanophoraceae), but it is not known if it was responsible for their deaths.

Environmental Requirements

In tropical regions (e.g., Papua New Guinea) A. novae-zelandiae appears to occur in cooler, high elevation habitats.


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Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year

Means of Movement and Dispersal

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Natural Dispersal

Like many other Armillaria species, A. novae-zelandiae spreads vegetatively from colonized woody matter by means of rhizomorphs. Infection of live hosts by this means leads to the formation of small disease centres.

There are strong indications that in New Zealand A. novae-zelandiae basidiospores (to a much greater extent than those of A. limonea) play a significant role in initiating new colonies, perhaps due to a better survival ability and plentiful rainfall throughout much of the year (Termorshuizen, 2000). Non-lethally infected trees were widely distributed in second rotation stands of P. radiata in a plantation forest originally established on a treeless plain (MacKenzie and Self, 1988). In the preceding first rotation stands the pathogen was present only saprotrophically (Gilmour, 1954), implying invasion by spores of dead trees and woody matter, giving rise to inoculum that infected trees in the following crop. These observations were supported by the mapping of high densities of colonies (an average minimum of 44 per hectare of only Anovae-zelandiae) infecting the second rotation trees in the same forest(Hood and Sandberg, 1993a; Hood et al., 2002c). Similar results were found in some kiwifruit orchards where individual colonies of Anovae-zelandiae were confined to single or several contiguously grouped, herbicide-treated willow stumps from which the pathogen had spread vegetatively along root systems to infect the immediately adjacent kiwifruit vines (Horner, 1992). Using molecular techniques, Dodd et al. (2010) identified A. novae-zelandiae as the predominant species among isolates from both kiwifruit orchards and second and third rotation P. radiata stands. High densities of coloniesof both A. novae-zelandiae and A. limonea, were mapped in indigenous forests and in first rotation Pradiata stands that replaced them after clearing and burning the original cover (Benjamin and Newhook, 1984a; Hood and Sandberg, 1987; 1989; 1993b). However, any new colonies detected after burning were only of A. novae-zelandiae.

Other studies have demonstrated the ability of artificially applied or naturally trapped basidiospores of Anovae-zelandiae to colonise partly buried billets and, to a lesser extent, stumps of Pradiata (Hood et al., 2002a; 2008; Hood and Gardner, 2005). Basidiospores of A. novae-zelandiae, identified using PCR, were readily detected using an air-sampling suction trap up to 150 m from the edge of a podocarp-hardwood forest during periods of fruiting (Power et al., 2008).

Accidental Introduction

Low to no risk of accidental introduction - see “Risk of Introduction” section.

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Forestry Yes Yes
Horticulture Yes Yes
Nursery trade Yes Yes

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Plants or parts of plants Yes Yes
Wind Yes

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Growing medium accompanying plants Yes Pest or symptoms usually visible to the naked eye
Roots Yes Yes Pest or symptoms usually visible to the naked eye
Seedlings/Micropropagated plants Yes Yes Pest or symptoms usually visible to the naked eye
Stems (above ground)/Shoots/Trunks/Branches Yes Yes Pest or symptoms usually visible to the naked eye

Impact Summary

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Cultural/amenity Negative
Economic/livelihood Negative

Economic Impact

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General remarks on the significance of A. novae-zelandiae in different crops and regions can be found in the section “Notes on Crops/Other Plants Affected”. This section focuses primarily on the effect of A. novae-zelandiae in New Zealand, as this is where the majority of the research on the impact of A. novae-zelandiae has been located. Growth loss and economic analyses in New Zealand Pradiata plantations have been undertaken by Shaw and Calderon (1977), MacKenzie (1987), Self and MacKenzie (1995) and Kimberley at al. (2002).

In New Zealand, Armillaria novae-zelandiae at one time caused serious damage in Pinus radiata plantations and in kiwifruit orchards. General accounts of the disease in forests have been published by Gilmour (1966), Shaw et al. (1976), Ridley and Dick (2001) and van der Pas et al. (2008), and reviews were produced by Hood (1989). Hill (1988; 2000), Hill and Travis (1994), Hill et al. (1999), Horner (1985; 1987; 1988a, b; 1990a, b; 1992; 2005a, b; 2006a, b) and Horner and Kay (2001) have discussed the disease in kiwifruit orchards.

First rotation pine plantations on cleared indigenous forest sites

Little indigenous forest is now converted to exotic pine plantation in New Zealand, but this practice was widespread during the 1970s and 1980s leading to severe mortality due to Armillaria (A. novae-zelandiae and A. limonea) in the young pine crops (Beveridge et al., 1973). Tracts of remnant podocarp-hardwood and southern beech forest previously logged of commercially valuable trees were clear felled, desiccated and burnt prior to planting. Heat from the fire temporarily reduced the level of Armillaria inoculum (Hood and Sandberg, 1989), but recolonization of dead wood was rapid, with signs of mycelial fans, rhizomorphs and (in season) prolific fruiting by A. novae-zelandiae (and A. limonea) on charred stumps and logs within 1-2 years. Mortality of pine seedlings commenced 3 to 6 months after planting and continued at an increasing rate for 3-5 years before declining (MacKenzie and Shaw, 1977; Roth et al., 1979; van der Pas, 1981a). Resultant mortality gaps were associated with indigenous hardwood stump root systems, which acted as a substrate providing an enhanced inoculum potential (Shaw and Calderon, 1977; Roth et al., 1979; van der Pas, 1981b; Benjamin 1983; Benjamin and Newhook, 1984b; van der Pas and Hood, 1984; Hood and Sandberg, 1993b). Mortality then declined up to about age 10 years, but infection continued through the remainder of the 25- to 30-year rotation in approximately 15% of the residual crop trees in a cryptic, non-lethal form (trees infected at the root collar retained green crowns and appeared healthy). There was no evidence of secondary spread between pine trees (Shaw and Toes, 1977; Roth et al., 1979; MacKenzie, 1987).

Estimates of seedling deaths during the first 5 years ranged 10-50% over an area of between 50,000 and 60,000 ha of land formerly covered in remnant podocarp-hardwood forest, with lower values on former southern beech forest sites (Shaw and Calderon, 1977; van der Pas, 1981a; Hood, 1989). The uneven distribution of mortality resulted in potentially lower stem wood quality in surviving trees because of excessive branch growth and hence knot size in stand gaps. Estimates of wood volume loss per unit land area in a rotation varied between 6% and 32% as a result of a combination of underutilized ground space due to mortality gaps and reduced growth increment from non-lethal infection in surviving trees (Shaw and Calderon, 1977; MacKenzie, 1987; Kimberley at al., 2002).

Second rotation pine plantations

There is now little planting of pine directly onto indigenous forest cutover sites in New Zealand, and early mortality in second and third rotation pine stands on such sites is generally lower (up to 5% of trees killed in small, scattered disease centres) and of little concern, with some exceptions (van der Pas, 1981a; MacKenzie and Self, 1988). Non-lethal infection is widespread, but variable in pine stands throughout New Zealand, ranging from less than 1% to 38% of trees affected, depending on the pre-plantation vegetation cover (Hood and Sandberg, 1993a; Self et al., 1998; Hood et al., 2002c). Non-lethal infection makes up a significant but smaller component of the overall growth loss (Kimberley et al., 2002).

Similar low early mortality and significant subsequent non-lethal infection occurs in plantations on originally non-forested land that have replaced first rotation stands in which the pathogen was not present, except as a saprotroph colonizing thinning stumps and dead trees, probably introduced by means of spore inoculum (Gilmour, 1954; MacKenzie and Self, 1988; Hood and Sandberg, 1993a; Hood et al., 2002c). Growth loss in one such stand was estimated as a little over 2% at 13 years (mid rotation; Kimberley et al., 2002).

Occasional, more severe mortality has occurred in pine stands subjected to atypical management procedures. For example, the long-abandoned practice of poison thinning, as the dead trees provide plentiful pathogen substrate for increased inoculum potential (van der Pas, 1981a). Establishing the stand after, for instance, a Eucalyptus crop produces a planting site containing hardwood instead of softwood pine stumps, which is more favorable to the fungus. Spraying with treated sewage effluent in stands on sites set aside for the disposal of such material  makes the trees stressed and more vulnerable to infection, due either to excessive water loading or to undesirable chemicals. Trees infected by Armillaria sp. Show a greater reduction in growth increment if also infected by the needle pathogen Dothistroma septosporum (Shaw and Toes, 1977) or if pruned too heavily.

Kiwifruit orchards

Armillaria novae-zelandiae had a serious impact in orchards of kiwifruit in parts of New Zealand during the 1980s and 1990s (Cutler and Hill, 1994). Spores colonizing stumps created during the removal of certain rows of the shelter belt species, Salix matsudana, to reduce shade as trees grew, enabled infection to spread into kiwifruit vines through extensive root interaction, resulting in sizeable mortality centres (Horner, 1992). Due to remedial measures the disease is no longer considered to be of economic significance in orchards of this host (Kiwifruit Vine Health, 2019a).

Impact: Biodiversity

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Armillaria novae-zelandiae is not known to threaten species or affect biodiversity.

Social Impact

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In parks and urban gardens in New Zealand, A. novae-zelandiae is most common in introduced hosts, and is responsible for a significant number of deaths of amenity trees and shrubs. For instance, it has been responsible for the loss of some trees in the Christchurch Botanic Gardens in the South Island of New Zealand (Bradbury and Steinegg, 1999). Similar issues in botanical gardens in Hobart and Melbourne were caused by Aluteobuballina.

Risk and Impact Factors

Top of page Invasiveness
  • Invasive in its native range
  • Has a broad native range
  • Abundant in its native range
  • Pioneering in disturbed areas
  • Tolerant of shade
  • Capable of securing and ingesting a wide range of food
  • Has high genetic variability
Impact outcomes
  • Host damage
  • Negatively impacts agriculture
  • Negatively impacts forestry
  • Negatively impacts livelihoods
  • Negatively impacts animal/plant collections
Impact mechanisms
  • Pest and disease transmission
  • Parasitism (incl. parasitoid)
  • Pathogenic
Likelihood of entry/control
  • Difficult/costly to control


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Economic Value

Armillaria novae-zelandiae has no economic benefit. In New Zealand there was at one time some interest in the association of A. novae-zelandiae with Gastrodia orchid rhizomes with the possibility of potential herbal medicinal use in China, but no industry has developed.

Social Benefit

Young fruitbodies of A. novae-zelandiae are edible. Known as “harore” they are harvested by some Maori.

Uses List

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Human food and beverage

  • Food


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To identify the species first requires laboratory isolation of fungal cultures from infected tissue, mycelial sheeting, rhizomorphs or, if present, fruitbody tissue. This is done by antiseptically transferring small pieces onto plates of an Armillaria-selective medium such as malt extract agar with suitable antibiotic &/or biocidal additives (e.g., o-phenylphenol or benomyl) to discourage contaminating bacteria and other unwanted fungi (Morrison et al., 1991; Worrall, 1991). One recipe prescribes the addition of from 2 to 10 ppm benomyl and (after autoclaving and cooling before pouring plates) 100 ppm streptomycin sulphate to a 2% malt extract agar base. A substitute may need to be found if the fungicide Benlateä (active ingredient 50% benomyl) becomes unavailable. Cultures may be maintained in the laboratory by sub-culturing on potato dextrose or 2% malt extract agar. Armillaria isolates are readily recognized in culture from their reddish-purple crustose appearance and production of characteristic rhizomorphs (distinct in appearance, however, from those formed in the field), but additional techniques are necessary for distinguishing species.

An older procedure to identify an unknown diploid field isolate, first developed with Northern Hemisphere Armillaria species (Korhonen, 1978), was to pair it with a range of selected, white, fluffy, single spore, haploid, tester isolates of known species. The paired tester isolate that changed from white fluffy to a dark purple, crustose, diploid form identified the unknown as being of the same species (Kile and Watling, 1988; Hood and Sandberg, 1987). The method is mostly reliable but takes time and the result is not always clear cut. Isolates of A. novae-zelandiae can also be distinguished from those of A. limonea by the production of rhizomorphs only in the latter after growing for approximately three weeks under a 24-hour photoperiod using specified lighting conditions (Shaw et al., 1981; Benjamin, 1983; Hood and Sandberg, 1987).

More recently, cultures of A. novae-zelandiae have been distinguished from other Armillaria species using molecular techniques (Coetzee et al., 2001b; 2003a; Maphosa et al., 2006; Pildain et al., 2009; 2010; Ota et al., 2011). Molecular procedures have been developed for identifying A. novae-zelandiae using the internal transcribed spacer ITS1 and ITS4 regions and the 5.8S ribosomal gene in Australia (Dunne et al., 2002; Smith-White et al., 2002) and New Zealand (Dodd et al., 2006; 2010).

In New Zealand rhizomorphs of the two common species are difficult to distinguish in the field (Hood and Sandberg, 1987) but according to Benjamin (1983) those of A. novae-zelandiae are slightly thicker on average and produced in greater abundance than those of A. limonea.

Detection and Inspection

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As with other Armillaria species, first evidence of disease in a crop will be symptoms of wilting or crown discoloration. Removal of soil to expose the root collar using a small trowel or grubber may reveal evidence of oozing resin or gum in conifers or hardwood species, respectively. This presence of resinosis or gummosis indicates damage to the living host, implying that the pathogen is acting parasitically and not simply feeding saprotrophically on already dead tissue. Black, branching, bootlace-like rhizomorphs consisting of a thin, outer, black rind and a soft, inner, pinkish, hyphal core, may be present. Also defining will be characteristic, thick, white mycelial sheets, fans or ribbons which are revealed by using a knife to cut and prize away the bark. If the host is already dead and no longer resisting the pathogen, these sheets may be present more extensively running up into the stem. Rarely, fruitbodies may be present (see “Description”, above). Unlikely in crop plants, but in older, mature trees, a butt heartwood rot may be present with a distinctive moist, cheesy, yellowish, fibrous decay interspersed with thin, black zone lines (the edges of sclerotial sheeting forming large, 2-3 cm “pockets” within the wood) which give rise to a scalloped appearance on drying.

Similarities to Other Species/Conditions

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Armillaria novae-zelandiae is one of many Armillaria species broadly similar in appearance and behavior (Watling et al., 1991; Pegler, 2000). All have white basidiospores, an anulus or ring (partial veil) around the fruitbody stipe (toadstool stalk), and many produce rhizomorphs (species without a ring belong in the recently erected genus, Desarmillaria in Koch et al., 2017). Species vary in their level of intrinsic pathogenicity against different hosts, but many are responsible for economic diseases in crops, plantations and managed natural forests around the world (Kile et al., 1991; Hood et al., 1991). All species also feed saprotrophically, colonizing and producing a white rot in wood, on which they fruit. Molecular analysis has shown that A. novae-zelandiae belongs in a broad group of Southern Hemisphere Armillaria species that cluster separately from those in the Northern Hemisphere (Maphosa et al., 2006; Coetzee et al., 2011; Klopfenstein et al., 2017).

Unusually for Armillaria species, A. novae-zelandiae was first recognized as a distinct taxon on morphological grounds (Stevenson, 1964). However, to distinguish it confidently from other species internationally requires cultural or molecular procedures (see “Diagnosis”, below).

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Research was undertaken in New Zealand from the late 1990s to develop an economic integrated control procedure for A. novae-zelandiae in second and third rotation Pinus radiata plantations as an alternative to the costly removal of previous crop stumps prior to planting, which though effective was generally not implemented operationally (see below). This included investigating host genetic resistance or tolerance, the effect of stock quality and vigour, potential biological control and a possible silvicultural management approach. Research in intensively managed kiwifruit orchards in the 1980s led to successful control of the disease. In both forest plantations and kiwifruit orchards in New Zealand, A. novae-zelandiae is no longer considered to cause a serious economic disease, although sporadic, locally important outbreaks may occasionally still occur in forests.

Cultural Control and Sanitary Measures

Planting at a greater density has been recommended on infested sites in Pinus radiata plantations, to compensate for mortality losses. However, this may only be effective if mortality gaps are not large and surviving trees are reasonably evenly distributed (MacKenzie and Self, 1988). In a trial in a second rotation Pradiata stand set up to determine how thinning influences the level of disease on infested sites, mean non-lethal infection was present in 50% of trees after a first thinning at age 7 years to 250 stems per hectare at age 13 years (approximately mid-rotation). This compared with 35% for the same treatment but with stumps simultaneously extracted, and 29-32% for unthinned controls (Hood et al., 2002c). It appears that the additional substrate provided by the thinning stumps had increased the inoculum potential (approximately half of the stumps were colonized by Armillaria). However, by age 19 years no difference was detected, apparently because of the increased vigour and associated physiological resistance of the host trees in response to thinning (Hood and Kimberley, 2009). Although there was a small loss in individual tree growth due to infection during the first half of the rotation, overall reduction for the full rotation was negligible. It was concluded that although thinning was not detrimental to the final crop volume, it gave no benefit in terms of disease control.

Field experiments have demonstrated the importance of the quality of the planting stock on sites known to be infested with A. novae-zelandiae (and A. limonea). In a first rotation Pradiata stand on a site cleared of podocarp-hardwood forest in New Zealand mortality from the disease among newly planted seedlings was lower than among rooted cuttings (Klomp and Hong, 1985). However, with improved quality of Pradiata cuttings, the trend was reversed when a second rotation trial stand was established on the same site (Hood et al., 2006). In each case, better planting stock was at an advantage when exposed to Armillaria inoculum.

Physical/Mechanical Control

Removal of stumps and woody debris before planting is the most effective method of controlling armillaria root disease. However, this procedure has been applied only occasionally in forest plantations because benefits are not seen by forest managers to outweigh the initial expense, despite economic analyses to the contrary (Shaw and Calderon, 1977; MacKenzie, 1987; Self and MacKenzie, 1995). Unlike some other management options, stump extraction also benefits subsequent rotations by removing inoculum from the site, although its effects on soil quality (removal of topsoil, compaction) must also be considered. Soil cultivation without stump removal has a number of advantages e.g., improved planting efficiency and better establishment, but it is not clear if this procedure reduces disease levels.

In one study, removal of indigenous forest stumps reduced mortality among planted pine seedlings to 12-21% after 5 years, compared with 52% on the untreated site (Shaw and Calderon, 1977; van der Pas, 1981a). In another, similar trial, which included clearing debris into windrows after removing indigenous forest stumps, mortality after 4 years was 1% in treated and 23% in untreated plots (van der Pas and Hood, 1984). In a third trial, in a second rotation radiata pine plantation following Pinus ponderosa that had been planted on a cleared indigenous forest site and subsequently poison thinned, mortality was reduced after 5 years from 22% (untreated) to 5% (stumps removed) at one site and from 10% to less than 1% at another (Self and MacKenzie, 1995). After 8 years, non-lethal infection in the same trial was reduced from 67% to 31% of trees affected and from 85% to 10% of trees, in the respective sites.

With smaller pine root systems, whole tree harvesting is another possibility prior to establishing second and third rotation stands that has not been investigated.

Various techniques were tried in infested kiwifruit orchards during the 1980s in New Zealand, including inserting root barriers consisting of trenches lined with polyvinyl chloride sheeting to prevent spread (Horner, 1987; Hill, 1988). However, the most effective procedure was using machinery to excavate and mechanically sieve the soil free from roots and other woody matter before replanting. This was undertaken operationally and achieved eradication of the pathogen in more than 90% of treated sites. At that time the high value of the crop justified the intensity and expense, but it was important to ensure that the ground area sifted encompassed the full extent of the infected root systems for full effectiveness. In a later study to investigate treatment of infected vines without having to remove and replace them, exposing root collars by water sluicing in order to surgically detach diseased tissue achieved significant disease control, and proved better than fungicide injection or application of biological control agents (Horner, 2006a, b). Final recommendations for kiwifruit vines were to clear stumps and woody matter from soil before establishing new orchards, to deploy shelterbelts of tree species with non-spreading root systems and to plant them at final stocking in order to eliminate the need for felling and creating stumps (Hill, 1988; Horner, 2005b). If shelterbelts had to be removed, stumps should be extracted and not sprayed with herbicides, which aided spore colonization by Anovae-zelandiae.

Biological Control

Research was conducted in the 1980s to evaluate the effectiveness of strains of Trichoderma species in controlling armillaria root disease in kiwifruit orchards and Pinus radiata plantations. In one trial pine seedling root systems were immersed in a selected Trichoderma slurry treatment before planting on an Armillaria-infested site. Mortality after 2 years was 6% of treated plants compared with 22% in the untreated controls (Cutler and Hill, 1994). Surviving treated plants were healthier. Similar results have been obtained in forest nurseries (Robert Hill, personal communication; Dyck, 2006).

Laboratory studies conducted with different Trichoderma species (e.g., T. atroviride, Tharzianum, T. viride) identified a number of potentially superior isolates in terms of their aggressiveness towards A. novae-zelandiae (Taylor, 1991; Cutler and Hill, 1994; Dodd-Wilson, 1996). Formulations were prepared, using substrates such as sawdust and peat, for testing in field trials (Hill and Travis, 1994), and some products were registered commercially for operational use in kiwifruit orchards (Cutler and Hill, 1994). Further tests with various formulations, e.g., as pastes and by injecting vines, have given variable results (Hill and Travis, 1994; McLaughlin, 1996; Hunt, 1998; Hunt and Clarkin, 1998; Horner 2006a, b). Trichoderma products are currently used in kiwifruit orchards against other pathogens (particularly Pseudomonas syringae pv actinidiae) and for general vine health (Woodcock, 2016; Kiwifruit Vine Health, 2019b).

In interactive laboratory studies with wood decay fungi as potential biological control agents for treating stumps, there was evidence of reduced colonization by A. novae-zelandiae of woody substrate also inoculated with either Rigidoporus concrescens or a species of Ganoderma (Li and Hood, 1992). However, in a field trial there was no evidence that these and other basidiomycete wood decay species reduced the level of A. novae-zelandiae when inoculated into Pradiata thinning stumps (Hood et al., 2015).

Chemical Control

Studies have been undertaken to test the effectiveness of chemical treatments in reducing the incidence of disease caused by A. novae-zelandiae (and A. limonea), although none have been used operationally in forest plantations. Mortality 4 years after planting P. radiata on an indigenous forest cutover site was 9% in plots with stumps treated with a commercial hydrocarbon mixture containing methyl isothiocyanate compared with 23% in untreated plots, despite there being no difference in the frequencies and quantities of soil rhizomorphs between sites (van der Pas and Hood, 1984). A similar result was obtained after applying agricultural lime to the soil surface. Sodium pentachlorphenate and pentachlorphenol introduced to the soil gave no protection to container grown P. radiata against artificial inoculum of Anovae-zelandiae (Shaw et al., 1980).

Soil fumigation was tried experimentally for controlling the disease in kiwifruit orchards using several chemicals, such as methyl bromide, chloropicrin, and dazomet (Ian Horner, pers. comm.). Methyl bromide was the most effective, but infected kiwifruit vine tap roots were found to grow to depths of 5 m, beyond the reach of the fumigants. In another study, treatment with a phosphorous acid product proved to be ineffective (Ian Horner, personal communication).

Host Resistance (Incl. Vaccination)

There is evidence that host species vary in resistance or tolerance to A. novae-zelandiae. In inoculation studies, Benjamin and Newhook (1984b) found more seedlings of P. radiata became infected and died than did those of Eucalyptus species, among which variation also occurred. Field observations of the effects of armillaria root disease (A. novae-zelandiae and A. limonea) on tree species planted on sites cleared of indigenous forest have been published (Weston, 1957; Gilmour, 1966). Most susceptible were P. radiata, species of Larix and Chamaecyparislawsonianaarl., while least affected were Cryptomeria japonica and Thuja plicata (Hocking and Mayfield, 1939; Jolliffe, 1940; Lysaght, 1942). Douglas fir (Pseudotsuga menziesii) was as susceptible as P. radiata but showed lower rates of mortality. Losses were low among eucalypts (Newhook, 1964). Although apparently less vulnerable, C. japonica was found to develop butt heart rot caused by Armillaria. A list of species recorded as susceptible to the disease in New Zealand (without distinguishing between Anovae-zelandiae and A. limonea) was published by van der Pas et al. (2008).

In inoculation experiments, no differences were found in infection and mortality among 25 Pradiata clones (Hood et al., 2009). More testing may eventually identify stock that is less susceptible, but with little more than a century’s exposure to a new pathogen (P. radiata is exotic in New Zealand) there may have been insufficient time or opportunity for resistant material of this host to become naturally selected. Fruiting is not common in P. radiata plantation forests in which the establishment of new disease centres may be from external spore sources, exposing trees to only part of the population gene pool.

Monitoring and Surveillance (Incl. Remote Sensing)

Monitoring of mortality is normally undertaken on the ground by means of transects or other appropriate field design (Hood et al., 2002b). In P. radiata plantations, non-lethal infection is estimated in the same way, except that in the absence of crown symptoms it is also necessary to expose the root collar to reveal the presence and extent of girdling infection (Shaw and Toes, 1977). Firth and Brownlie (2002) found that by using high resolution colour, stereo aerial photography it was possible to detect most trees greater than 2 m tall that had been killed by Armillaria, reducing the time and investment necessary for ground monitoring. Mapping of infested sites in second rotation stands indicated that the distribution of non-lethally infected trees showed a broad approximation to that of trees visibly killed by the disease (e.g., areas with an average pre-thinning mortality greater than 3% of trees killed were reasonable indicators of stand areas with non-lethal infection exceeding 25%). This implied that it might be possible to use the distribution of visible mortality (e.g., from aerial photography) to produce contour maps that predict and delineate areas requiring treatment, either during or at the end of the rotation (Hood et al., 2002b; Hood et al., 2006).


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16/10/19 Original text by:

Ian Hood, New Zealand Forest Research Institute (Scion), Rotorua, New Zealand.

Charles Shaw, Consultant, USA

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