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Virus Goodwill from BioStim

With all the bad news in the world, I want to do something positive. As a local small business, we are feeling the strain as well. If you have received the government stimulus or have a few extra dollars, we hope to entice you to make a purchase.

For the next week, we will include a 20 gram sample of MycoGold with every order. We hope you will give it to someone who might benefit (drop in their letter box and message them) or pass it onto a family member to use.

This is just our little goodwill effort to help get our thoughts off the virus. Get into the garden/land and grow a plant. The vitamin D and fresh air can only be beneficial 🙂

p.s If you have a business and would like some free promotion, send me a reply email with your details to feature in our next newsletter.  

Kind regards
Tim Lester

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By the way, what actually are ‘saprophytes’?

Saprophytic plants, literally, are plants that live of rotting material (sapros = rotting, and phyton = plant in Greek), but in fact, no plant have been found yet which can use dead organic material for food directly.
Anyway, these plants have no chlorophyll in their cells, which means they are unable to assimilate carbon by themselves. They have no green leaves, often they even have no leaves at all. Saprophytes are mostly whitish, but can have brightly coloured flowers. They grow in places with lots of rotting dead leaves, often in deep shade in tropical forests.

In their underground parts (rhizomes or roots) are certain cells that are filled with structures (hyphae) of soil fungi. Often, but not always, these fungi are capable of ‘digesting’ the rotting material and converting it by enzymes into molecules (sugar) which they can feed on. So, the fungi are the real saprophytes, living of rotting material. Now, the plants without chlorophyll digest the fungus that live inside their roots or rhizomes, thus they are not autotrophic/self supporting, but heterotrophic plants (hetero = another, trophein = feed). And because they are living on fungi they are called myco-heterotrophic plants / MHP’s (mycos = fungus). This mycorrhiza (mycos = fungus, rhizon = root) of MHP’s makes it possible for them to grow in places with not enough light for ordinary autotrophic plants to survive. The same might be the case for places without enough nutrients in the soil.

To complicate matters there is evidence that some fungi neither are saprophytes but have underground connections with big forest trees or other autotrophic plants. So the trees, the fungi, and the myco-heterotrophic plants all three together form a kind of plant community, a symbiosis (living together), to make it possible for the MHP to live. In the special case of MHP’s, the linking fungus delivers the assimilated carbon from the autotrophic plant to the myco-heterotrophic plant.

We are still very much interested in new collections of saps, and we are always willing to identify them.
Our knowledge about saprophytes from Africa and Asia is less extensive, but we are interested to study them as well (especially in the genus Thismia).

Here some hints, how to collect saprophytes:

  • If you find one it is likely that there are more, since circumstances seem to be favorable for this mode of life.
  • Most important is to preserve specimens in spirit (roots, buds, flowers, fruits).
  • Do not forget to collect the root system if possible.
  • Look for pollinators, smell, and something about the flower biology
  • Make drawings or take colour slides.
In general: take some time to have a good look at the plants when you find them: they deserve it!
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Achlorophyllous plants

Achlorophyllous plants are puzzling! They are always characterized by remarkable reductions concerning root, shoot and leaf structure and sometimes even hardly resemble a flowering plant. It is no wonder that they are often collected by mycologists! Here some basic information:

Since assimilation of carbohydrates through photosynthesis without chlorophyll is impossible (as far as we know) and the direct metabolization of dead organic material has never been detected in flowering plants, achlorophyllous plants must have another source of carbon. They split up in two distinct groups:


These plants develop special organs (haustoria) penetrating foreign plant tissue in order to participate at least to some extend from their host’s assimilates (carbon compounds), water or nutrient uptake (e.g. Dodder, Broomrapes).

Mycoheterotrophic Plants (‘Saprophytes’)

In these plants a fungus (or several?) lives inside their roots (‘mycorrhiza’) providing all requirements for plants growth. This is the plant group that I investigate.

Voyria truncata just emerged upon the soil surface. The pencil-sharpener serves as a scale

More than 400 species, in 87 genera and 11 families, of mycoheterotrophic plants have been described. Of those, only orchids and members of the Monotropaceae (Indian-Pipe Family) are fairly well investigated. Information about the other families are scarce. The most recent work on the neglected genera has been done by Hiltje and Paul Maas and co-workers in Utrecht/Netherlands. Nevertheless, root structures (morphology, anatomy, mycorrhiza) are often entirely unknown. Most probably this is due to the remote and hardly accessible habitat of these plants, the deep shaded tropical rainforest, and the fact that they are easily overlooked in the field (see the picture to your left!). Hence, they get found by mycologists!

Lately, I focused on the genus Voyria of the Gentianaceae (Gentian Family) where 19 species have been distinguished so far, all except one living in tropical America. Then I looked after Triuridaceae and Burmanniaceae (TriurisSciaphilaBurmanniaDictyostega). Momentarily, I’m working on Burmanniaceae and Polygalaceae (AfrothismiaEpirixanthes). All of these plants share some morphological characters with Voyria but are not at all related to them. I could show that their mycorrhiza is an arbuscular mycorrhiza (AM), a form of fugus-plant-symbiosis which is very well known for more than hundred years. However, the achlorophyllous species so far investigated revealed some very unique features, yet unknown despite the long and intensive research on AM.

My approach was led by the following questions. Answers I found so far are indicated shortly:

What kind of morphological/anatomical adaptations have evolved in connection to its special life form?
At least one of those adaptations is a ‘condensation’ of the root system (becoming short and thick).
Are mycorrhizas in myco-heterotrophic species different from mycorrhizas in autotrophic species?
Yes, definitely in Voyria tenella, V. obconica, V. aphylla, Triuris hyalina, and Afrothismia winkleri, less pronounced but still different in Voyira truncata, Burmannia tenella, and Dictyostega orobanchoides. More strange mycorrhizal patterns may be anticipated.
Do the mycorrhizas between various myco-heterotrophic species differ?
Yes they do, only in Voyria tenella and V. obconica I found the same ‘intraradical fungus garden’.
What do mycorrhizal structures tell us about taxonomy and systematics?
The closely related Voyria tenella and V. obconica do have the same mycorrhiza whereas V. aphylla shows an intermediate pattern, linking to the mycorrhiza of V. truncata and the autotrophic gentians. The two Burmanniaceae Burmannia tenella and Dictyostega orobanchoides show at least in the root cortices the same intracellular hyphal pattern. Afrothismia winkleri, a Burmanniaceae from Africa, however, has an entirely deviating mycorrhizal pattern (although it is an AM!).
How do these plants use their root fungus?
Very sophisticated!! Please read the abstracts e.g. on Afrothismia winkleri and Voyria tenella .
What is the actual carbon source?
From the observed direct hyphal bridges between roots of neighboring plants and the achlorophyllous plants we must infer the carbon (and most probably everything else too) must come from the neighboring plant.
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Orchids and Fungi

by Ian St George

Sentences like “In vesicular-orbuscular mycorrhizas, the mycobiont undergoes pronounced alterations in morphogenesis involving appressoriuin formation, (and) arbuscule development…” leave the amateur gasping for breath. This is a complex subject with its own expert language: fortunately writers like Warcup and Perkins from Australia demonstrate the ability to convey complex information simply.


What is a fungus? We think of toadstools, bracket fungi, ringworm, athlete’s foot – they are all fungi. The fungi are called Mycota, about 50,000 species described, including mushrooms, yeasts, rusts, smuts, mildews and molds. The mushroom we eat is the fruit (also known as the “perfect state”) of a larger organism, which has hollow branched filaments called hyphae that form networks in the soil; the bracket fungus we see is the fruit of a larger organism whose hyphae thread the dead bark and wood of the tree. Ringworm is the infection of the human skin by similar organisms; if you scrape the skin cells and look at them under the microscope you can see fungal hyphae.

The study of fungi is mycology, and many words referring to fungi have “myc-” in them. Mycoses are fungal diseases. ascomycetes are a genus of fungi, Mycetophilidae are the fungus – loving gnats. Medical mycologists identify pathogenic fungi by their asexual spores; plant mycologists may deal with fungi that develop perfect states – the fruit that contain sexual spores – and use these for identification. Where perfect states cannot be achieved in cultivation, the structure of the hyphae, the pattern of branching and rejoining of hyphae (anastomosis), or the number of nuclei in cells may give a clue as to identity.


If you read about orchids you can’t escape phrases like “fungal associations”, and “pine needle layer rich in fungal hyphae”. In fact the roots of most vascular plants have evolved in association with soil fungi. The resulting combined structures are called mycorrhizas (“fungus-roots”). There are seven main kinds of mycorrhizas, the four most carefully studied involving crop plants, forest trees, heaths and orchids [1].

Many orchid mycorrhizal fungi belong to the form genera RhizoctoniaEpulorrhiza, or Ceratorhiza. These same genera may contain species that are orchid pathogens, form associations with other plants, or have no plant associations.

The habitat of the fungus may determine the habitat of the orchid – thus the fungus Rhizoctonia borealis requires acid soils under conifers, so that is where its associated European orchids Spiranthes gracilis and Goodyera repens are found.

Strictly, the term mycorrhiza should apply only to the fungus/root association, but it is loosely association between the fungus and the developing orchid protocorm (the stage between seed and embryo).

Orchid mycorrhizas

Orchids require the relationship with a fungus for their existence. The importance differs among species, the “infection” by the fungus being heaviest in temperate terrestrials, but light in tropical epiphytes. The relationship is essential for the germination of the seed of all orchids in the wild, and remains essential for a few species throughout life.

Seeds, protocorms and fungi

Orchid seeds are tiny and lack the built-in nutrition of bigger seeds; orchids then pass through a nongreen (“achlorophyllous”) developmental stage when they cannot use fats, break down starch, obtain phosphates or photosynthesise, and therefore rely on an external source. This is provided either by man in the form of simple carbon-containing foods in sterile seed germination, or by a fungus which breaks down complex compounds into simpler ones in symbiotic germination. The fungal hyphae penetrate via the base end of the seed. The hyphae enter the cells and coil into structures called pelotons Germination of the seed into a protocorm follows. The cells eventually digest the pelotons, but occasionally the fungi become parasitic and destroy the protocorm.

Roots and fungi

In some species (GastrodiaDanhatchia and Corybas cryptanthus in New Zealand) chlorophyll never does develop, so the orchids rely for all their lives on associations with fungi. In others, the leaf-size is too small to support the rest of the orchid, and the orchid continues to rely partly on the fungus for its nutrition (Corybas cheesemanii for instance); such plants have been called saprophytic, but that is an incorrect application of the term. Some plants of the European Spiranthes spiralis pass alternate seasons underground, apparently fully nourished by their fungus during that time; some NZ orchids do not appear above ground every year and may do the same.

Most terrestrials seem to thrive better in the wild than in pots (some cannot be cultivated “artificially” at all), probably because they must have access to at least some of their nutrition via their ~ association.

In different terrestrial orchids the fungi penetrate the stems, tubers or root hairs, via epidermal (“skin”) cells after hyphae have spread over the root surface [2]. Pelotons are formed, and eventually digested.

“Symbiosis” suggests mutual benefit and indeed Cymbidium and its fungus each require the vitamin thiamine, made up of thiazole and pyridine; the fungus supplies the thiazole and the orchid supplies the pyridine [3]. Most orchid-associated fungi can, however, live without the orchid, and it seems that whereas the fungus supplies the orchid with a range of nutrients and stimuli, the orchid usually provides little in return.

Many orchids have “host” cells that store fungus, and adjacent digestion cells that break the fungus down by means of substances known as phytoalexins. The partnership between orchid and fungus has been called symbiosis (a ‘Swan situation’ as the politicians say in Wellington these days), or a “delicately balanced mutual antagonism’ [4], or plain parasitism (of the orchid on the fungus, that is.

Fungi that are apparently symbiotic can turn nasty and attack the orchid; furthermore the fungi of epiphytes may invade the orchid’s host tree to the tree’s (and ultimately the orchid’s) detriment.


Some studies in the laboratory suggest that specific orchids require specific fungi, but few associations have been studied in the wild. Fungi are difficult to isolate and difficult to grow (especially to the usually readily identifiable perfect state), and even in one orchid species, the fungus required by the protocorm may be different from that required by the adult. Certainly some orchids can establish successful relations with several different fungi.

Perkins has looked at the Australian orchids Pterostylis acuminata and Microtis parviflora in the wild and in the laboratory [19, 20]. Whereas only a few species of fungi were associated in the wild, several more would form associations in the laboratory – thus, ecological specificity” (what happens in the wild) is different from “potential specificity” (what could happen if laboratory experiments were to reflect the wild state) [18].

New Zealand Studies

In 1911 Lancaster showed that fungal hyphae do penetrate the root hairs of NZ epiphytes and form pelotons which are digested by the orchid cells [5].

Ella Campbell began a series of papers on the fungal associations of NZ’s nongreen orchids in 1962 [6-10]; she showed: –

  • Gastrodia cunninghamii is associated with the fungus Armililaria mellea which is itself a parasite on the roots of forest trees [6].
  • G. minor is associated with and derives nutrients from an unidentified fungus which also penetrates the roots of adjacent manuka [7].
  • What is probably the bracket fungus Fomes mastopourus inhabits the roots of Acacia melanoxylon and is an endophyte of Gastrodia aff. sesamoides, which digests it [8];

All around the roots of taraire trees grow the hyphae of the puftball fungus Lycoperdon perelatum, and these form a network around and attach to the rhizomes of Danhatchia australis, invade the tips of root hairs, and are digested by the cells of the orchid. The orchid is parasitic on the fungus, which in turn derives nutrients from, and may damage, the roots of the taraire [9].

Corybas cryptanthus has an associated unidentified fungus that invades the roots through root hairs attached at tiny conical projections; the fungus spreads among the beech leaf litter, and is a weak parasite on the Nothofagus[10].

Australian work

J.H. Warcup, M. Clements, K. Dixon and A. Perkins and their co-workers have been the major contributors to the study of Australian orchid/fungus relationships [2, 11-21]. Readers interested in delving deeper are referred to these authors (for instance Warcup [16] gives an excellent general overview of the fungal relationships of South Australian orchids). Here are a few snippets.

  • Warcup and Talbot seem to have had a genius for inducing orchid fungi to fruit in culture. They grew fungi from pelotons teased from the cells of Australian native orchids – of 102 isolates from 25 orchid species 66 fungi were induced to fruit. Fungal species of the following genera formed mycorrhizal associations with orchid:: species (of the genera in brackets): Thanatophorous(AcianthusThelymitra), Ceratobasidium (PterostyisPrasophyllumAcianthus), Tulasnella (DiurisAcianthusThelymitraCaladeniaCymbidiumDendrobium) and Sebacina (AcianthusCaladeniaGlossodiaMicrotis); the same fungal species often formed mycorrhizal associations with European orchid species. These truly intracellular fungi were often different from those found on the surface of orchid roots [12].
  • Fire affected the abundance, behaviour and composition of fungus infecting West Australian orchids; there were six categories of fungus, and each was specific and consistent within species and within most genera; the rare and related DrakaeaParacaleana and Spiculaea had a unique and culturally distinct fungus noted for its intense violet-pink colour [2].
  • Initial contact between fungus and seed is haphazard – there is no evidence that an attractant is used by the orchid seed. Seeds appeared to resist entry by incompatible fungi, while allowing the entry of compatible fungi. There was a strong specificity of fungus for each orchid studied. Pelotons appeared about a week after initial infection in some cells and signified a compatible orchid/fungus match that would lead to germination. The protocorm seemed to have entry, holding and digestion zones for the fungus, though the way the fungus is controlled in these zones is unknown. Failure of germination was caused by fungal hyphae failing to penetrate the seed, or by penetrating all the embryo’s cells resulting in death of the embryo [21].
  • In Pterostylis the fungi can be grouped, and where the groups are found is determined by the environment. One fungus, for instance, was found only under Pinus radiata. Geographic distribution (and perhaps some aspects of habitus?) of orchid species may thus be determined by fungal ecology [17].
  • Perkins and co-workers found only two fungi associated with Microtis parviflora in the wild, and the same two in protocorms: they concluded that the adult roots associate with a narrow range of fungi in the field (have a narrow ecological specificity) and these assmiations are established in the protocom. On the other hand, many fungi were able to form associations with M. parviflora in the laboratory, indicating a broad potential specificity [19].
  • It would seem logical that the germination of orchid seed in the wild should depend on the amount of fungus in the soil, but this may not be so. Perkins and co-workers studied Pterostylis acuminata and its fungal associations: this orchid appears to associate with only one specific fungus, a subspecies of Rhizoctonia solani. Furthermore this orchid reproduces asexually (i.e. essentially by cloning). The orchid and the fungus may therefore be co-distributed, and if an orchid is able to establish at a new site, the resultant increase in the associated fungus may favour further spread of the orchid. There are implications here for the resiting of rare orchids – if there is a single fungus associated with the orchid, a new site will need to be apt for the fungus as well as for the orchid: if the fungus does not survive, neither will the orchid [20].

Orchids that form ecologically specific relationships with single pollinating insects can only survive in the presence of that specific insect. We now see that there are orchids which form ecologically specific relationships with single mycorrhizal fungi: they can only survive in the presence of that specific fungus. How these observations apply to the New Zealand species is open to speculation.


I am grateful to Dr A.J. Perkins for supplying a list of Australasian papers.


  1. Peterson RL, Farquhar ML. Mycorrhizas – integrated development between roots and fungi. Mycologia 1994; 86 (3): 311-326.
  2. Ramsay RR, Dixon KW, Sivasithamparam K. Patterns of infection and endophyte associations with Western Australian orchids. Lindleyana 1986; 1: 203-214.
  3. Hijner JA, Arditti J. Orchid mycorrhiza; vitamin requirements and production by the symbionts. Amer.J.Bot. 1973; 60: 829-835.
  4. Arditti J, Fundamentals of orchid biology. Wiley, New York, 1992. p445.
  5. Lancaster T.L. Preliminary note on the fungi of the New Zealand epiphytic orchids. Trans.N.Z.L 1911; 43: 186-191.
  6. Campbell E.O. The mycorrhiza of Gastrodia cunninghamii HookF. Trans.Roy.Soc.N.Z 1962; Bot 1: 289.
  7. Campbell E.O. Gastrodia minor Petrie, an epiparasite of Manuka. Trans.Roy.Soc.N.Z 1963; Bat 2: 73.
  8. Campbell E.O. The fungal association of a colony of Gastrodia sesamoides R.Br. Trans.Roy.Soc.N.Z. 1964; Bot 2: 237.
  9. Campbell E.O. The Fungal Association of Yoania australisTrans.Roy.Soc.N.Z. 1970; Biol.Sci. 12: 5-12.
  10. Campbell E.O. The Morphology of the Fungal Association of Corybas cryptanthusJ.Roy.Soc.N.Z 1972; 2: 43-47.
  11. Dixon K. Seeder/clonal concepts in Western Australian orchids. In Population ecology of terrestrial orchids. Eds T.C.E. Wells and J.H. Willems. J.H. SPB Academic Publishing: The Hague, 1991, ppl11-124.
  12. Warcup J.H. and Talbot P.H.B. Perfect states of Rhizoctonias associated with orchids I-III. New Phytologist 1967; 66: 631-641; 1971; 70: pp35-40; 1980; 86: pp267-272.
  13. Warcup J. H. Specificity of mycorrhiza association in some Australian orchids. New Phytologist 1971; 70: pp41-46.
  14. Warcup J. H. Symbiotic germination of some Australian orchids. New Phytologist 1973; 72: pp387-392.
  15. Warcup J.H. ne mycorrhizal relationship of Australian orchids. New Phytologist 1981; 87: pp371-387.
  16. Warcup J. H. Mycorrhiza. In Orchids of South Australia. Eds R.J. Bates and J.Z. Weber. Flora and Fauna of South Australia Handbook Committee, Adelaide, 1990. pp21-26.
  17. Ramsay R.R. Sivasithamparam K. and Dixon K.W. Anastomosis groups among Rhizoctonia-like endophytic fungi in south western Australian Pterostyis species. Lindleana 1987; 2: pp161-167.
  18. Matsuhara G. and Katsuya K. In situ and in vitro specificity between Rhizoctonia spp. And Spiranthes sinensisNew Phytol. 1994; 127: 711-718.
  19. Perkins A.J. Masuhara G. McGee P.A. Specificity of the associations between Microtis parviflora and its mycorrhizal fungi Australian J. Bot. 1995; 43: pp85-91.
  20. Perkins A.J. and McGee P.A Distribution of the orchid mycorhiza1 fungus, Rhizoctonia solani, in relation to its host Pterostyis acuminata, in the field. Australian Journal of Botany 1995; 3(6): pp565-575.
  21. Clements M. Orchid mycorrhizal associations. Lindleyana 1988; 3: pp73-86.
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The Myth of Foliar Feeding

“Fertilizers sprayed on the leaves of trees and shrubs are more effective than soil applications?”


By Linda Chalker-Scott, Ph.D., Extension Horticulturist and Associate Professor, Puyallup Research and Extension Center, Washington State University


Recently, I received an email from a professional colleague whose clients often ask about foliar feeding
as a method of fertilizing plants. As he says, “All the water soluble fertilizer companies advertise the
practice all the time.” What, he wondered, was my opinion of the practice?

Foliar feeding involves spraying the foliage of target plants with water-based fertilizers. The logic for the
practice is based on scientific research from the 1950’s, which demonstrated that leaves can take up
minerals through their stomata, and in some cases through their cuticles. This research is consistently
cited in the argument that foliar feeding is 8, 10, or even 20 times more effective than traditional soil

In assessing the advertised claims for foliar feeding of shrubs and trees, I had particular questions that are
answered in bulleted lists below (all bullets are directly from marketing media):

(1) What are the advantages of foliar feeding over soil application?
• Immediate results
• Prolong bloom
• Increase crop yields
• Increase storage life of food crops
• Boost growth during dry spells
• Increase cold and heat tolerance
• Increase pest and disease resistance
• Maximize plant health and quality
• Help the internal circulation of the plant


(2) When should one use foliar nutrients sprays?
• When the soil is too cold for conversion of nutrient elements into usable forms
• When it is at least 72°F
• Any time except when it is too hot or too cold
• Transplant time
• Bloom time
• When a quick growth response is desired
• After fruit set
• Every 2-3 weeks
• Any time of stress
• As long as the plant has leaves that aren’t dormant
• When the soil is deficient in nutrients


(3) What time of the day, and in what quantity, should you apply foliar fertilizers?
• Early morning
• Until it drips from the leaves
• There is no improper way


(4) How long will material last on the leaves?
• 24 hours
• 1-2 days
• Four weeks

(5) What nutrients are critical components of foliar feed fertilizers?
• Nitrogen
• Phosphorus
• Micronutrients

(6) Apart from commercial formulas, what should homemade mixtures contain?
• Seaweed
• Compost tea
• Natural apple cider vinegar
• Blackstrap molasses
• Fish emulsion
• Baking soda
As one company states, “In our opinion, foliar feeding is by far the best approach to use to insure
maximum growth, yields, and quality by overcoming limitations of the soil and its ability to transfer
nutrients into the plant.”

The Reality

If these laundry lists look more like a multiple choice test rather than solid information, it’s not surprising.
Foliar feeding is yet another agricultural practice best suited to intensive crop production under specific
soil limitations rather than as a landscape management tool. Thus, advertisers take great liberties with the
facts, often resulting in contradictory messages (note especially the recommended temperature
conditions!). Rather than individually refute the numerous errors in the claims, I’ll explain when foliar
feeding might actually be beneficial.

The original 1950’s research came from Michigan State University and was particularly useful in
understanding how nutrients move within plant tissues. As explained by Dr. Tukey in his testimony to
the Joint Committee on Atomic Energy, use of radiolabelled nutrients allowed his team to discover
“…that a leaf is a very efficient organ of absorption. The amounts may at first seem relatively small, but
to offset this handicap, the efficiency is high.” From this advertisers claim that foliar feeding is 8, 10 or
20 times more effective than soil application. This is not accurate for several reasons.

Obviously, materials applied directly to a leaf are more likely to enter the leaf in large quantity than the
same materials applied to the soil. Leaching, chemical reactions, microbial activity, etc. can decrease
what actually reaches the roots and is taken up into the plant. But materials applied to the leaf do not
necessarily travel throughout the entire plant as effectively as they do through root uptake. They often
remain in the same or adjoining tissues but travel no further. This is especially true of those elements
recognized as “immobile” within plant tissues (apart from root uptake and xylem transport).

Research over many decades has explored the mineral uptake and transport of many species of fruit trees,
conifers including pine and spruce species, and some hardwoods of ornamental or commercial value.
Results have been mixed in many cases, with some species responding well to treatment and others
remaining unaffected. Generally, the results suggest that foliar application of particular nutrients can be
useful in crop production situations where soil conditions limit nutrient availability.


For instance, alkaline soils do not readily release many metallic nutrients, especially iron and manganese. Zinc,
copper, magnesium, molybdenum, boron, and calcium are other micronutrients required in small
quantities that have been applied to foliage in an effort to relieve deficiencies and combat fruit disorders.
Fruit, as adjacent tissue, can benefit from foliar spray. But this is a localized application that does not
affect the trunk or roots – and therefore is not a solution to soil imbalances. In fact, researchers
consistently state that foliar treatments are a specialized, temporary solution to leaf and fruit deficiencies
in tree fruit production but will not solve larger soil management issues.

On the other hand, macronutrients, such as nitrogen, phosphorus and potassium, are needed in larger
quantities. While many of these are mobile in the plant, it is pointless to apply them to foliage as leaves
cannot take up enough material to supply the entire plant’s demands. Furthermore, foliar application of
high concentrations of such nutrients often results in leaf burn as water evaporates and the fertilizer salts
remain behind. Substituting numerous, lower concentration applications would not be cost effective.

Species differ widely in their ability to take up nutrients through their leaves. Differences in cuticle
thickness, stomatal resistance, and other genetic factors will influence uptake, as will environmental
conditions. Plants in a protected situation (like a greenhouse) have thinner and more porous cuticles than
plants in the field and take up foliar sprays much more readily. Likewise, plants adapted to arid
environments naturally have thicker, less penetrable cuticles than those from more moderate locations.

A better management solution to the problem of nutrient availability is to choose plants that can adapt to
the existing soil conditions. If you have alkaline or calcareous soils, for heaven’s sake don’t install acid
loving plants! Poor plant selection in terms of mineral nutrition will be a management problem for the
lifetime of the plant – which may be pretty short. Choose cultivars of species that are more resistant to
alkaline soils – they are able to acidify the root environment so that micronutrients are remobilized from
the soil and available for uptake.

The existing research does not justify foliar fertilization of landscape plants as a general method of
mineral nutrition. It can be useful for diagnosing deficiencies; for instance, spraying leaves with iron
chelate can help determine if interveinal chlorosis is from iron deficiency. It would obviously have
benefit for those landowners with landscape fruit trees that perpetually have flower or fruit disorders
associated with micronutrient deficiencies. Applying fertilizers to leaves (or the soil) without regard to
actual mineral needs wastes time and money, can injure plant roots and soil organisms, and contributes to
the increasing problem of environmental pollution. 


The Bottom Line
• Tree and shrub species differ dramatically in their ability to absorb foliar fertilizers.
• Proper plant selection relative to soil type is crucial to appropriate mineral nutrition.
• Foliar spraying is best accomplished on overcast, cool days to reduce leaf burn.
• In landscape plants, foliar spraying can test for nutrient deficiencies, but not solve them.
• Micronutrients are the only minerals that are effectively applied through foliar application.
• Foliar application will not alleviate mineral deficiencies in roots or subsequent crown growth.
• Foliar spraying is only a temporary solution to the larger problem of soil nutrient availability.
• Minerals (especially micronutrients) applied in amounts that exceed a plant’s needs can injure or
kill the plant and contribute to environmental pollution.
• Any benefit from foliar spraying of landscape trees and shrubs is minor considering the cost and
labor required.