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Uptake preservation of CUT FLOWERS & FOLIAGE A report for the Rural Industries Research and Development Corporation By Margaret Johnston, Alison Fuss, Helen Murphy, Nely Moncada, Daryl Joyce and Bhesh Bhandari. April 2000 RIRDC Publication No. 00/66 RIRDC Project No. UQ-53A

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© 2000 Rural Industries Research and Development Corportaion. All rights reserved. ISBN 0 642 58096 0 ISSN 1440-6845 Uptake preservation of cut flowers and foliage Publication No. 00/66 Project No. UQ-53A The views expressed and the conclusions reached in this publication are those of the author and not necessarily those of persons consulted. RIRDC shall not be responsible in any way whatsoever to any person who relies in whole or in part on the contents of this report. This report is copyright. However, RIRDC encourages wide dissemination of its research, providing the Corporation is clearly acknowledged. For any other enquiries concerning reproduction, contact the Publications Manager on phone 02 6272 3186. Researcher Contact Details Dr Margaret Johnston Senior Lecturer The University of Queensland Gatton GATTON QLD 4343 Phone: 07 5460 1240 Fax: 07 5460 1455 Email: M.Johnston@mailbox.uq.edu.au Website: http://www.aghort.uq.edu.au RIRDC Contact Details Rural Industries Research and Development Corporation Level 1, AMA House 42 Macquarie Street BARTON ACT 2600 PO Box 4776 KINGSTON ACT 2604 Phone: 02 6272 4539 Fax: 02 6272 5877 Email: rirdc@rirdc.gov.au Website: http://www.rirdc.gov.au Published in June 2000 Printed on environmentally friendly paper by Canprint

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Foreword Drying and dyeing cutflowers and foliage is an important way of adding value. This project was undertaken to meet the industry’s needs for more reliable and cost-effective methods of preserving flowers and foliage. The report provides useful insights and technical information of value to the industry. This report, a new addition to RIRDC’s diverse range of over 450 research publications, forms part of our New Plants Products R&D program, which aims to facilitate the development of new industries based on plants or plant products that have commercial potential for Australia. This project was funded from RIRDC Core Funds which are provided by the Federal Government and is an addition to RIRDC’s diverse range of over 450 research publications. It forms part of our Wildflowers and Native Plants R&D program, which aims to improve the profitability, productivity and sustainability of the Australian wildflower and native plant industry. Most of our publications are available for viewing, downloading or purchasing online through our website: • downloads at www.rirdc.gov.au/reports/Index.htm • purchases at www.rirdc.gov.au/eshop Peter Core Managing Director Rural Industries Research and Development Corporation

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Acknowledgements This project was funded by Rural Industries Research and Development Corporation. The authors thank: • Mr and Mrs David and Olive Hockings for providing fresh and preserved Stenanthemum stems; • Mr and Mrs Graham and Esther Cook for providing fresh riceflower stems; • Mr Geoff Sullivan and the members of AUSBUD for advice and providing eucalypt

stems; • Mr Alan Lisle for assistance with statistical analysis.

About the Authors • Dr Margaret Johnston Senior Lecturer, The University of Queensland Gatton has been involved

with floricultural research for over 20 years with UQ Gatton and Department of Primary Industries, Queensland. Her particular research interest is the problems associated with the domestication of Australian native plant species and she has developed koala fern (Caustis blakei).

• Dr Alison Fuss, Horticultural Consultant, has a research background in the development of

Australian native plants for the ornamental industry. She has assisted with the writing up of this project.

• Dr Helen Murphy, Research Officer, The University of Queensland Gatton has recently completed

her PhD on Guar and has been assisting with the completion of this project. • Ms Nely Moncada was enrolled in a Research Masters on the topic ‘Uptake preservation of cut

flowers and foliage of Australian native plants’ and conducted the experimental work for this project.

• Professor Daryl Joyce is a Postharvest Scientist and originally the Principal Investigator until he

resigned from The University of Queensland Gatton and accepted a professorial appointment with Silsoe College, Cranfield University, UK.

• Dr Bhesh Bhandari Lecturer, The University of Queensland Gatton, has been involved with food

dehydration, water activity and microencapsulation research. He has been with The University of Queensland since 1993.

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Contents

Foreword ................................................................................................................................. iii

Acknowledgements ................................................................................................................. iv

About the Authors.................................................................................................................... iv

Executive Summary ................................................................................................................ vi

1. What are the current industry practices and problems in the preservation of cut flowers and foliages in Australia? .............................................................................1

2. How does the anatomy and morphology of plants influence the uptake of preservative solutions?...................................................................................................8

3. Do postharvest environmental factors affect the uptake of glycerol solution by Eucalyptus robusta? ...................................................................................................16

4. Is the uptake of preservative solution by Eucalyptus robusta and E. tetragona affected by the type of humectant used and its concentration?.................37

5. Do adjuvants affect the uptake of glycerol solution by Eucalyptus robusta and E. tetragona?.............................................................................................................50

6. Can fluorescent dyes be used to characterise the uptake of glycerol solution by Eucalyptus robusta?.........................................................................................................61

7. Do plant growth regulators affect the uptake of glycerol solution by Eucalyptus robusta?.........................................................................................................68

8. Does the uptake of glycerol solution affect the respiration of cut Eucalyptus robusta and E. tetragona? .............................................................................79

9. Is the uptake of preservative solution by stenanthemum affected by the type of humectant used and its concentration? ..........................................................88

10. Does relative humidity and type of packaging during storage effect the outturn quality of glycerined stenanthemum cut flowers?.........96

11. Is the uptake of preservative solution by riceflower affected by the type of humectant used and its concentration?......................................................................103

12. What type of dye is most effective at colouring riceflower during uptake preservation?......................................................................................................108

13. Conclusions and industry implications ...........................................................................114

14. Bibliography....................................................................................................................117

15.Appendices......................................................................................................................118 Appendix 1: Information that is available on preserving cut flowers and foliage and the factors affecting it ...........................................................................................................118 Appendix 2: Colour photographs of anatomical characteristics......................................124 Appendix 3: Statistical results pertaining to Chapter 2 ...................................................128

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Executive Summary The participants of the initial industry workshops conducted in Queensland and Western Australia agreed that research was needed to address the key problems faced by the preserving industry in Australia. Hence this project was designed using a systematic scientific approach to identify appropriate chemicals and to optimise protocols for the uptake preservation of a selection of native cut flowers and foliages. The importance of fresh and healthy plant tissues for uptake preservation was shown. Stems to be preserved by uptake should be handled in a similar way to those stems to be sold as fresh cut flowers. The time between harvest and preservation needs to be minimised (less than 12 h) and stems should be recut before being placed into preservative solutions. Prewilting offers no advantage over processing material immediately after harvesting. The uptake of preservative solutions can be enhanced by modifying the prevailing environmental conditions during transportation, handling and processing. Appropriate conditions could result in more rapid and uniform solution uptake. There are some very simple procedures and equipment, which could be put in place to improve the outturn quality of stems preserved by uptake preservation and in turn its economic return. Continuous elevated lighting with 36 W cool white fluorescent lamps, to give 56-75 µEinm-2 s-1 should be installed approximately 25 cm above the foliage being processed. These lights should remain illuminated throughout the uptake process. Air circulation at low airspeeds can increase the uptake of glycerol solution by cut stems. High airspeeds can be detrimental to the uptake of solution. Hence, fans should be installed in the processing area and operated on low during processing. The uptake of glycerol solution by cut stems is improved by increasing the vapour pressure difference between the leaf and the atmosphere. This can be achieved by increasing the temperature during processing. Uptake is very poor at or below 7°C. Glycerol was found to be an appropriate preservative solution for Eucalyptus foliage, and flowering stems of stenanthemum and riceflower. The most rapid accumulation of glycerol in stems of E. robusta and E. tetragona was achieved with a 20% glycerol solution. Uptake of glycerol solutions by E. robusta, stenanthemum and riceflower is reduced after 3 to 4 days, but this did not occur for E. tetragona. For both riceflower and stenanthemum, it is possible to reduce the time required to take up the required amount of glycerol by increasing the concentration of the glycerol solution used. Polyethylene glycol (PEG), at the concentration and in the combination with glycerol tested, is an inappropriate humectant for the preservation of E. robusta and E. tetragona, as it caused discoloration and loss of flexibility of the foliage. It is unclear whether PEG alone or in combination with glycerol can be taken up by stenanthemum to a suitable level to provide any preservation of quality of the plant material. However, PEG does not seem to be a suitable humectant treatment for riceflower as uptake of solutions containing PEG was very slow. Pyranine dye can be used to assist in studying the movement of glycerol solution through cut stems of E. robusta. However, the degree of fluorescence caused by pyranine dye cannot be used as a measure of the amount of solution taken up by cut stems of E. robusta, unless it is standardised for each solution type. Based on the location of fluorescence in the stem and leaf tissues, glycerol moves relatively quickly and is distributed fairly evenly throughout these tissues over the initial 3 day period. In fact after as little as 24 hours of uptake of glycerol solution, the solution had been distributed to the peripheral leaves. The commercial adjuvants, Agrimul PG 2067 and Geropon SDS, at the concentration tested (100 mg mL-1), should not be added to a glycerol solution during uptake preservation of cut stems of either E. robusta or E. tetragona. The addition of Agrimul at this concentration does not provide any benefits and the addition of Geropon SDS was detrimental to the stems.

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Further experimentation is required to determine whether the use of adjuvants can provide any benefit to the uptake preservation process for E. robusta and E. tetragona. Such experimentation should consider the use of Agrimul PG 2067 and Geropon SDS at lower concentrations, as well as other adjuvants, like Agral 600. Testing needs to be done on a species by species basis. The plant growth regulators, ABA and fusicoccin, at the concentrations tested, should not be used to pre-treat cut stems of E. robusta for uptake preservation with glycerol solution. These plant growth regulators do not provide any benefits. Further experimentation is required to determine whether the use of these or other plant growth regulators may provide any benefit to the uptake preservation process for E. robusta. Such experimentation should consider varying the duration of the pulse, varying the concentration of the plant growth regulators in the pulsing or uptake solution, varying the application technique, (ie direct application to the foliage as a spray or as a dip), as well as using alternative chemicals that control stomatal aperture. Moreover, testing may need to be done on a species by species basis. Red food dyes, added to uptake preservative solutions, may provide an appropriate, effective and efficient means of colouring the foliage and florets of riceflower. Further experimentation is required to determine if other coloured food dyes are also effective at colouring the foliage and florets of riceflower. Research is needed to determine the appropriate concentration and time of treatment to achieve a range of colour intensities. Textile dyes and Dutch dyes are inappropriate for use with uptake preservation solutions as a way of colouring the foliage and florets of riceflower, but may be effective during immersion dyeing. Processors need to consider the environmental conditions during the preservation process and storage to ensure outturn quality of preserved stems is achieved and maintained. Storage of preserved flowering stems of at high relative humidity (>80%) and low relative humidity (30%) should be avoided as quality will be adversely affected by excessive water adsorption by humectants or by moisture loss respectively. If stems are to be packed for prolonged storage, cartons should have a low density polyethylene (LDPE) liner and a sachet of glucose or potassium chloride to maintain a stable environment and hence avoid loss of outturn quality. Adoption of the equilibrium relative humidity (ERH) test could determine the suitability of a product to the environmental conditions in the consumers display area. Alternatively it could be used as a feedback loop to refine the preservation process. The research conducted during this project has provide the dried and dye sectors of the Floricultural Industry with some useable results which can be incorporate simply into their own processing operations. Minor modifications to current practices can add to the efficiencies of the process, significantly improve outturn quality of these flowers and foliages and should ultimately result in better economic returns for the industry. The importance of using high quality products that were handled appropriately after harvest, as well as improving the environmental conditions during processing and understanding the different requirement for different species is of great importance.

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1. What are the current industry practices and problems in the preservation of cut flowers and foliages in Australia?

Introduction Whilst most cut flowers and foliages exported from Australia are fresh, there is an expanding industry based around the drying and preserving of cut flower and foliages. In 1992-93, Australia’s dried flower exports (including preserved material) were valued at A$ 6.246 million (Karingal Consultants, 1994). Nearly all of the flowers and foliages that are dried or preserved are Australian native plant species, with much of the production coming out of Western Australia. Recently there has been an expansion of the drying and preserving industry in Queensland, with the development of commercial plantations of its indigenous species, including eucalypts, riceflower and stenanthemum. To date there has been very limited public research undertaken to optimise the protocols and identify appropriate chemicals for preservation of native cut flowers and foliages. Individuals in the industry have developed many of the processes used to dry and preserve these species, but it has usually been undertaken on a non-systematic, trial and error basis. Thus, despite the considerable economic importance of this industry it remains poorly defined in terms of its products, processes and organisation. It is important that efforts be made to develop improved processing technology to support this growing industry and maximise the outturn quality of its products. In order to achieve these improvements, it is essential to firstly define the current situation, in terms of products and processes, and identify the problems and difficulties faced by the industry. This chapter attempts to do this for the industries in Queensland and Western Australia.

Objectives 1. Define the current products and processes used by the industry in Queensland and Western

Australia to preserve native cut flowers and foliages. 2. Identify the problems and difficulties faced by the native cut flower and foliage preserving

industry in Queensland and Western Australia.

Methodology Face to face workshops were held with industry groups in Queensland and Western Australia. The workshops in Western Australia were held in Perth and Albany on 9 and 10 April 1996, respectively. These workshops were conducted by Dr Margaret Johnston and Mr Mark Webb. The workshop in Queensland was held at the University of Queensland Gatton on 30 May 1996. This workshop was run by Dr Daryl Joyce, Dr Margaret Johnston, Dr Bhesh Bhandari and Ms Nely Moncada. The workshops were fairly informal, with a list of questions used as a program to ensure that all relevant topics were covered (Table 1.1).

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Table 1.1 List of questions used at each of the industry workshops held in 1996 to gather appropriate information on preservation of flowers and foliages.

1. What are the chemicals currently being used to preserve flowers and foliages?

2. What is the ideal treatment for preserving flowers and foliages?

- What are the characteristics of the chemicals?

- What are the characteristics of the processes?

3. How are the flowers and foliages handled prior to preservation?

- During harvesting and transport?

- During processing?

- During storage and handling?

4. What factors cause variability in the quality of preserved flowers and foliages?

a. season/environment

b. agromonic factors

c. maturity

d. genotype

e. processing variables (postharvest environment)

f. pest (diseases/decay)

g. packaging/bleeding

5. Based on industry experience, clearly state the key problems with preserved flowers and foliages. (The intention is to use this list of problems to define experimental treatment limits).

6. What are the industry priorities in relation to preserving flowers and foliages?

Detailed results Summary of Western Australian workshops The workshop held in Perth attracted 29 people and that in Albany was attended by 11 industry representatives. The industry representatives at these workshops felt that the major problems facing the preserving industry related to the poor postharvest handling of plant material harvested from natural bush and the length of time taken for these products to get to the processing plant. Most of the processing operations are based in Perth, while much of the product is harvested in the lower south-west of Western Australia. Thus, it can take from 1 to 3 days to transport the products before they are processed. Many of the smaller processors commented that they had few problems when they picked the product locally and placed it into the preservative solution that day. Peter Ashby, of Dakota Enterprises (a supplier of a preservative solution), agreed that better results could be achieved if farmers who harvested wildflowers on their own property treated the product immediately. Most of the processors had no environmental control in the sheds where they undertook the preserving. Thus they found that while they could successfully preserve a small quantity of the product, it was often much harder to get reliable and consistent results when preserving large volumes of product.

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The lack of specialisation in the preservation industry was considered to be a problem. All processors tried to preserve all products instead of specialising in a few lines and preserving those very well. Notes on the various products presented later in this report indicate the degree of diversity with which the industry has to deal. The industry representatives had observed that some products, particularly those that were slow growing, were more difficult to preserve. The major problems that were reported included leaf drop, bleeding of the preservative solutions from the plant material and growth of moulds on preserved product. There was considerable discussion about the relative advantages of the uptake and immersion methods of preservation. The major disadvantage of the immersion method is that the stem as well as the foliage is dyed. However, the immersion method can be used to treat product that is too old for successful absorption. Although reluctant to give specific details, most of the industry members were using glycerol (30%) as the uptake preserving solution but some used a mixture of glycerol (20%) and polyethylene glycol (10%). They believed that this combination of solutions reduced the problem of bleeding. It was generally expected that during the preservation process a bunch would take up between 240-260mL of preserving solution (specific bunch size was not identified). The industry members were unwilling in public to discuss the chemicals that they were using; however some did reveal this individually. The anti-microbial compounds being used were sodium benzoate (0.1-0.2% by weight) or quaternary ammonium compounds. It was suggested by John Zucon, of Polymer Craft, that some of the anti-microbial compounds used by the cosmetic industry could be useful (orthophenol phenate) but they are costly and require a fairly high pH to be effective. Monoethylene glycol had some anti-fungal properties. Citric acid was being used to reduce pH to 4-4.5. The dyes used depended upon availability and cost. The red dye most commonly used was Tartrazine (ICI), and the blue dyes used were Edicol Blue (FCB) or Hid Acid Blue (German dye). It is well known that humectants, like glycerol, absorb water at high humidity and this may result in bleeding. It is understood that Darotech (a commercial company of the Department of Agriculture, Victoria) has patented a method of reducing bleeding of preserved plant material. It was suggested that this method involves the incorporation of a solid substance (10%) into the glycerol. Allegedly, glycerol will absorb 2-3 molecules of water while the substance will absorb up to 7 molecules of water, hence giving a 10% buffer against bleeding at high humidity. The industry representatives were keen to source additional information from the organisers of the workshops and the commercial suppliers of chemicals that were present. There were several queries regarding enhancing uptake of the preservative solutions. One grower said that bracken fern was being preserved in brine and sold for $3.50 per bunch wholesale. This was considered to be a good return for this weed species. The Western Australian Industry suggested the following species should be given priority in this study:

Agonis parviceps (fine/white ti tree) and/or Kunzea sp., (bottlebrush) Conospermum sp. (smoke bush), Bossiaea sp. (mini holly, waterbush) and Kauri hazel.

The industry members felt that success of preservation by the uptake method depended upon the length of time and environmental conditions to which the product was exposed between harvesting

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and arrival at the processing shed. They believed that it would be useful to have some experiments conducted in Western Australia. Comments about particular products processed in Western Australia Stirlingia latifolia (3.5 million stems preserved per annum) This product has a clearly defined season of 6-8 weeks in October and November. The stick-like stems are harvested from the wild, not for their flowers, but for the attractive display created by the fluffy husks that develop around the seeds. One problem is that Stirlingia is often picked too immature, before the fruit has developed into a full ball. This practice arises because bush-pickers are concerned that if they do not pick it that day someone else will. However, the industry representatives believed that Stirlingia was easier to preserve (i.e. takes up solution better) when picked at the correct stage. Stirlingia occurs locally in the Perth region and during its harvest season the processors work an evening shift to process the product bush-picked that day. The product is picked in the morning and delivered to the shed by 3 pm for processing which continues until 10 pm. Some members said that they held the product in a cold room in water but others said that the product is not placed into water prior to preserving. The industry representatives said they achieved an 80% success with preservation of this species and considered it to be “forgiving”. The industry representatives at the workshop in Albany said that there was very little Stirlingia processed as there was only a small amount available in the district. They experienced problems with fungal growth and fruit drop. Agonis spp. (Fine ti tree (A. parviceps) 3 million stems harvested per annum; Coarse ti tree (A. juniperina)) Three species of ti tree are processed: fine, coarse and Rosa. Most industry members reported that they had more problems with fine ti tree than the other species. The seasonal availability of each of the species varied. `Rosa’ flowered in summer (October to January), coarse ti tree flowered from summer to winter (February to June), while fine ti tree was only available in winter (June to October). The industry representatives believed that the problems they experience during processing ti tree were related to poor air circulation, particularly as the fine ti tree had many small leaves close together. They reported that when this product was placed into buckets, all leaves below the edge of the bucket fell off. By placing the stems into cups or a gutter system, they felt that there was improved air circulation around the stems and that the problem of leaf drop could be reduced. Some industry members had installed lights in their sheds and believed this had improved uptake of preserving solution. All industry representatives believed that many of the problems that they were experiencing related to the care of the product after bush-harvesting. This product was picked in the lower south west of Western Australia and placed into bundles containing 15 to 25 bunches. These bundles were then wrapped in plastic and transported on an enclosed but not refrigerated truck to Perth. They believed that it was 24-72 hrs between harvesting and processing and that the product was not given appropriate postharvest treatments during this time. In terms of rating the product for difficulty it was given a 3-4 out of a possible score of 10 (20-40% success rate). Often the leaves fell off 3 weeks after processing and the quality of the product 6 months after processing was poor. Industry representatives who attended the Albany workshop obtained the best results when the product was processed rapidly, ie. if it were placed into the preserving solution within 0.5-1 hour after harvesting and left for 24 hours. They used a gutter system rather than buckets and used fluorescent lights in the shed at night. Winter in Albany is cold and wet, and these conditions cause problems for processing.

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Kunzea ericafolia This has a fine lush green leaf and huge market acceptability. The use of this product for preserving was limited by leaf drop. It was rated as the most difficult species to preserve (unsuccessful). Davisia cordata This product is processed year round except during winter. It was rated as difficult to preserve (40% success rate) by all industry representatives. The major problem was leaf drop. Bossiaea sp. (Mini holly) This product was very difficult to preserve because of the leaf drop problem. The product is available all year, but industry representative said that they did not process it during the winter. Eucalyptus spp. This product could only be processed when it did not have soft tips which occur in late autumn and winter. Helichrysum cordatum (Sea Crest) This product was considered difficult to obtain, but preserves easily. Conospermum cassinervium (Tassel smoke) This is considered to be a very difficult product to preserve and received a rating of 4 on the scale of difficulty (only 20% success rate). Industry members believed that the time between harvesting and preserving was critical. They suggested that the product could be successfully preserved only if it were cut and placed into preservative solution immediately. Tassel smoke is slow growing, woody and has a thick cuticle. These characteristics were thought to contribute to its degree of difficulty in preservation. Adenanthos cuneata (Templetonia) Adenanthos cuneatus is available from spring to autumn. It processes well, but has soft tips in winter. Summary of Queensland workshop The industry members (25-30 people) listed a number of factors that they had noticed contributed to product viability. These included the time taken to get the harvested material into the preserving solution, the environmental conditions both in the field and in the preserving area and the maturity of the plant material used. They had noted that if Stenanthemum stems were placed into preserving solution within 3 hours of harvesting then the uptake of preserving solution was better than if it were left out of solution for a longer period. It was also reported that the cool moist conditions that prevail at Maleny seemed to cause less uptake of the preserving solution compared with warmer drier locations. Providing good ventilation or better air movement in the bucket had increased the uptake of the preserving solution by promoting a higher transpiration rate. An industry representative from Beerwah noted that the uptake of preservative solution from specially designed channels (viz. individual stems put through wires into the solution) was greater than from buckets. For Stenanthemum, the maturity of the plant material being processed also impacted on the quality of the finished product. It was suggested that removing immature foliage resulted in a better preserved product, as the young leaves were considered to serve as ‘sink’ for the uptake solution. As the solution builds up in the soft tissues, it apparently becomes toxic. Many different types of chemicals were being used for preserving plant material. For example, glycerol, ethylene glycol, PEG 400 and propylene glycol were used as humectants. Citric acid was often added to the uptake solution as it lowers the pH and thereby reduce microbial growth. A dip of

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Cislin and Rorvral may be used to eliminate insects and prevent fungal growth, respectively. Alginox is a preservative solution additive that can be used as a biocide in the uptake solution. Acetone has also been used in uptake solutions with a view to enhancing uptake. Bleaching chemicals have been used in uptake solutions to highlight veins and thus give an interesting effect. Dyes have been used similarly. The idea of supplying a perfume in the solution used for preservation was proposed. It was noted that there was a need for a relevant chemical database that would indicate a specific humectant for a particular plant material intended for a specific market. When the group was asked, “What constitutes the ideal treatment?”, the response indicated that only quality materials should be used. It was suggested that poor quality plant material should no longer be preserved and that the industry should set a standard for the quality of their product. If this could be achieved the group felt confident they would receive “the right price”. There was considered a need to educate consumers regarding the shelf life of the preserved product. Most felt that good quality preserved stems would last 3 to 6 months, and should command a good price in the market. There were some suggestions given for the need for better packaging. It was suggested that a fibre cardboard carton lined with polyethylene barrier facilitates better moisture exchange and maintains the quality of the preserved product. Some use a sachet of salt, which helps adjust the relative humidity inside the carton during transport and helps to maintain product quality. Several possible areas of research were identified. Market research into the consumers’ preference in terms of colour, use of scent, allergy issues, etc. was considered important for the growth of the industry. A comparison of commercial and laboratory practices (e.g. bunches in a bucket of solution) would be useful. It was considered that work was needed on the preservation of additional species; in particular, riceflower and kangaroo paw, as both of these species are grown extensively in Queensland. Riceflower is sometimes airdried and whilst it looks attractive it is extremely fragile. It would therefor be beneficial to develop suitable preservation technology. Similarly, kangaroo paw preserved with glycerol exhibits good quality. The concept of using treatment combinations: (e.g. uptake and immersion) needs to be explored.

Discussion of results The industry workshops conducted in Western Australia and Queensland indicated that the groups were experiencing very similar problems and difficulties in preserving flowers and foliages, despite the fact that they were working with quite different plant material. Not only were people from the two States concentrating on different plant species but also the sourcing of the material differed. In Western Australia much of the product used in preserving was sourced from natural bushland, while in Queensland product was predominantly harvested from cultivated stands. Variability in the preserved product was the main concern of the industry groups. While the natural variability in the product itself was identified as a problem, the time taken to get harvested material into the preserving solution and the lack of environmental controls during uptake preservation were also considered to be key factors contributing to this variability. The timing issue was exacerbated in Western Australia due to the large distances between the site of harvest and the processing facilities. It was suggested that this could extend up to 3 days with the problems being compounded by poor postharvest handling during that time. In both states, glycerol was the key preservative. It was used in solution either alone or in combination with other humectants, such as PEG. A range of anti-microbial chemicals and dyes were also added to the uptake solutions. Western Australia reported problems of leaf drop and the growth of moulds on the preserved product. It is likely that the long time between harvesting and preserving may have contributed to the leaf drop

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problem. It is probable that the ‘bleeding’ of glycerol from the leaves of the preserved material encourages the mould growth. Processors in Queensland were trying to overcome the development of moulds by dipping their product in a fungicide solution. The industry groups in both states were interested in the development of techniques that would control or stop the ‘bleeding’ particularly at high relative humidities and for specific markets. Overall, there was a general consensus that there was a lack of specialisation within the preserving industry in Australia. This was compounded by a lack of public research support in the area of optimising protocols and identifying appropriate chemicals for preservation of native cut flowers and foliages. To its credit the industry has achieved significant economic importance by its own invention and ingenuity. It now needs a systematic scientific approach to overcome the major hurdles it is facing. Growers commented that it was difficult to source information on drying and preserving and background information on the principles behind uptake preservation. To this end a list of references has been produced in Appendix 1.

Implications Research needs to address the key problems faced by the preserving industry in Australia. A systematic scientific approach is required to identify appropriate chemicals and to optimise protocols for the uptake preservation of native cut flowers and foliages.

Recommendations Experimentation in this project should concentrate on:

• Understanding the morphology of the plant material and how this influences preservation outturn quality,

• Identifying appropriate humectants, • Identifying chemicals which can improve the uptake of solution, • Developing techniques to minimise ‘bleeding’, • Determining the optimum time between harvesting and preserving, • Investigating the effect of prevailing environmental conditions, and • Assisting in the development of quality systems.

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2. How does the anatomy and morphology of plants influence the uptake of preservative solutions?

Introduction Uptake preservation is often used because it is the most simple and low cost method for small-scale preservation, and can be done at home or on the farm. Plant material preserved in this way often more closely resembles the natural appearance, than when processed by immersion. However, industry members have observed that “some species preserve easier and better than others”. The relationship between specific plant characteristics (such as vascular anatomy, stomatal size and distribution, cuticular wax and physiology), environmental conditions and humectant uptake are poorly defined (Joyce, 1998). This chapter aims to provide the reader with suitable background information to enhance their understanding and appreciation of the plant processes involved and exploited during uptake preservation.

Objectives 1. Provide background information on the anatomy and morphology of plants in relation to

uptake preservation.

2. Provide basic guidelines on the selection and handling of material for uptake preservation.

3. Record observed difference in anatomy and morphology of 4 Eucalyptus species and Stenanthenum scortechinii.

Brief Literature Review How do solutions move into and through plants? The ‘cohesion theory’ is used to explain uptake and movement of water and solutes through plants. It has three basic elements: ♦ the driving force resulting from the water potential gradient, ♦ the resistance or hydration forces that determine the interaction between water molecules and plant

tissue (adhesion), and ♦ the cohesion of water molecules (Salisbury and Ross, 1990). Water movement from the soil, through the plant and to the atmosphere is a continuum. Water moves from the soil, through the root hairs of the epidermis and through the root cortex to vascular tissues in the roots and stems. It then flows, via the vascular system, into the leaves, from where much water is lost by transpiration (evaporation) to the atmosphere. Loss of water from the leaves establishes the water potential gradient along the continuum. Anatomy and morphology of stem vascular tissue Water and solutes move through that part of the vascular tissue known as the xylem. The xylem forms a continuous capillary system from near the tip of the roots through the stem into the leaves, where it branches into a system of fine veins. The principal cell types in xylem are fibres, tracheids, vessels and parenchyma. Tracheids and vessels are dead cells, arranged contiguously (end to end) in order to conduct water, as well as to contribute to mechanical strength. They have pitted, pointed ends, matched to provide low resistance to water movement. The numerous vessel elements, which form the vessel members, have end walls that are perforated or completely lost to promote efficient water movement. Tracheids are smaller and thinner than vessels. Tracheids are typically about 10-25 µm (micrometre) in diameter,

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while vessel members are 40 to 80 µm, and rarely up to 500 µm (Salisbury and Ross, 1992). The other cells of the xylem, the parenchyma and the fibres, are involved in lateral transport, storage and support functions. Another tissue in the vascular bundle is the phloem. The cells in the phloem are sieve tube elements (with pitted sieve plates) and the dense, nucleated companion cells. This tissue is involved with the translocation of sugars and hormones around the plant. In woody plants, like Eucalyptus spp. and Stenanthenum scortechinii, secondary growth occurs at the vascular cambium giving rise to new xylem and phloem tissues each year. Large vessels are clearly obvious in transverse stem sections, and most water movement occurs through the newest of the vessels and tracheids (i.e. those less than 1 year old). The rate of fluid movement in the xylem is considered to be similar to water movement in small capillaries and is described by the modified Hagen-Poiseuille equation. This equation states that the movement of fluid is directly proportional to the water potential gradient between each end of the tube and to the radius of the tube (Hall, 1976; Salisbury and Ross, 1992; Gartner, 1995). The rate of flow is also inversely related to the length of the tube and to the viscosity of the solution. That is, as the radius of the vessel is doubled, the rate of flow is increased by 16-fold. Rapid flow of water through large vessels can, however, mean that there is a greater chance of cavitation occurring (Salisbury and Ross, 1992). That is, an air embolism or bubble may form comparatively easily within the vessel elements of the xylem. Anatomy and morphology of leaves Transpiration occurs primarily from leaves. Like stems and roots, leaves are composed of dermal, ground and vascular tissues. The dermal tissue consists of epidermal cells, trichomes and stomata, with their accessory and guard cells. The epidermal layer is covered by cuticular wax, which is a matrix of cutin, oligosaccharides and lipids (Martin and Juniper, 1970; Willmer, 1983; Taiz and Zeiger, 1991). The cuticle imparts a high resistance to diffusion of gases, including water vapour. Stomata are openings in the leaves for gaseous exchange. Trichomes (hairs), such as those (for example) which can be seen on the lower surface of Stenanthenum scortechinii leaves, decrease water loss by increasing boundary layer resistance (i.e. trapping water vapour in an unstirred boundary layer near the leaf) (Plate 2.1).

10

Plate 2.1 A section of a leaf of Stenanthenum scortechinii showing trichomes on the undersurface. Two layers of elongated palisade mesophyll can be seen beneath the epidermis, as well as spongy mesophyll in the lower part of the leaf, and the centrally located vascular bundle. The ground tissue of leaves is called the mesophyll. It usually has two layers, the upper palisade layer with elongated cells, and the spongy mesophyll layer with loosely packed irregularly shaped cells. The leaves of plants adapted to different environments show different amounts of these types of mesophyll. In plants adapted to a dry (xerophytic) habitat, palisade cells occur beneath both surfaces of the leaf (Esau, 1953). The vascular tissue in leaves forms a network of fine veins with a reticulate and/or parallel pattern. Following delivery via vascular tissues and adjacent ground tissue, water can be lost to the atmosphere from stomata, specialised hydathode pores and through the cuticle. Most water loss occurs via the stomata. When stomata are open, there is relatively little resistance to water vapour flow, and a high proportion of water is lost through them. When the stomata are closed, water may still be lost, albeit at a much reduced rate, from cracks and other small openings, ectodesmata and other pathways through the cuticle. In conifers and broad leaf trees, water loss from cuticular transpiration ranged from 2.5-3.0%, whereas stomatal transpiration accounted for 8.3-50% of the total water loss (Sebanek, 1992). The rate of stomatal transpiration is dependent more upon the localisation (exposure) of stomata and their aperture, rather than to their density (frequency). Water loss can even be quite high from partially closed stomata. The degree of wax impregnation of the cuticle is more important than the thickness of the cuticle in controlling water loss. Several authors have shown that the chemical composition of the impregnating wax markedly affects its efficiency as a water barrier (Kamp, 1930; Sitte and Rennier, 1963; Radleer 1965, all cited by Martin and Juniper 1970). However, Chambers and Possingham (1963) indicated that the longer and more complex the shape (form) of the wax platelets forming the wax layers, the more effective is their waterproofing capability.

11

What plant material should be used? Uptake of preserving solutions relies on transpiration, which occurs in living cells (Salisbury and Ross, 1992). Hence, it is important to use quality stems free of diseases and damage. Harvested stems need to be at the correct stage of development. Foliage with full sized leaves should be used as immature leaves wilt during the preserving process (Dubois and Joyce, 1990). How should material be handled in preparation for preserving? When cut stems are placed into water or preservative solution, the solution moves directly into the vascular tissue (xylem). However, if the stems are kept dry for long periods prior to preservation, air bubbles (embolisms) can form and impede the flow of water. Thus, cut stems should be placed directly into water and/or held in a cool environment. It is recommended that stems be recut under water immediately prior to standing them in the preservative solution (Dubois and Joyce, 1990). In general, principles applied to maintaining the vase life of cut flowers and foliage are applicable to preservation by uptake. There is no evidence that fraying or slitting of stems improves their uptake of preservation solution (Dubois and Joyce, 1992). Industry members have sometimes observed that uptake preservation is more successful when small batches are used. This is probably because overcrowding is avoided, and, hence, air circulation is relatively free. For such reasons, bunches need to be untied to assist uptake (Dubois and Joyce, 1990). Large quantities of cut stems in a shed will raise the relative humidity of the area due to the water loss to the atmosphere through transpiration. This may adversely affect uptake, unless good ventilation is maintained in the shed.

Methodology Four eucalypt species were used in this study: Eucalyptus robusta, E. tetragona, E. crucis and E. bakeri. Forty stems of E. robusta foliage were harvested from a managed garden at The University of Queensland, Gatton, while stems of the other three species, E. tetragona, E. crucis and E. bakeri, were harvested from a farm near Southbrook from 4 trees randomly selected within rows of each species. Stems were cut to 30 cm and held overnight at 5°C in buckets of deionised water covered with polyethylene sheets. Morphological characteristics of the leaves and stems of eucalypt species The morphological parameters were recorded for each of the four species including number of leaves, leaf thickness, leaf area, total leaf area and stem thickness from 20 replicate stems per species. Measurement of leaf area involved leaf removal hence 10 different stems were used. Total leaf area was calculated from mean number of leaves per stem and leaf area. Histological studies of the leaves and stems of four eucalypt species Light Microscopy Transverse leaf sections for each of the 4 eucalypt species were prepared for six replicate cut stems. The first fully expanded leaf plus the 1 cm long cylinders of stem tissue cut from the base of each 30 cm stem were fixed in formaldehyde acetic alcohol (90 : 5 : 5 of 70% ethanol : formaldehyde : acetic acid by vol.) for 24 hours. Fixed tissue was then dehydrated in a 70%, 85%, 95% and 100% tertiary butyl alcohol series. After dehydration, tissue was kept in a liquid paraffin : tertiary butyl alcohol (1:1, v/v) mixture for 30 minutes at 40°C, followed by 2 hours in Paraplast X-TPA paraffin wax (Monoject Scientific, Division of Sherwood Medical, St. Louis, U.S.A) at 60 °C, and then 24 hours in Paraplast PLUS paraffin wax at 60°C. The infiltrated tissue was then embedded in Paraplast PLUS. Sections were cut on an AO Spencer microtome. After dewaxing in three sequential 10 minutes changes of xylol and two changes of absolute ethanol for 1 minute, followed by 1 change of 70% ethanol and 1 change of deionised water, the sections were transferred to 1% aqueous Safranin O solution and stained for 3 minutes. Sections were then dehydrated through two 1 minute changes of each ethanol series- 70%, 90%, 100%, and counter stained in 1% absolute ethanol/clove oil based Fast Green for 15

12

seconds. Sections were cleared and permanently mounted in Xam neutral mounting medium (BDH, U.K.). The thickness of the upper and lower cuticle for six replicate transverse leaf sections was measured under light microscope with a calibrated eyepiece (1unit = 10µm). For each stem transverse section (n=6), the number of xylem vessels was recorded and the diameter of each xylem vessel was measured using a sampling protocol. Four sampling areas a, b, c and d were randomly chosen. Electron microscopy Leaf segments measuring about 1 mm2 were taken from the first fully expanded leaf of 4 stems for each of the 4 eucalypt species. Segments were fixed for 24 hours in 3% glutaraldehyde solution buffered to pH 6.8 with 0.1 mol/L sodium phosphate buffer. They were then rinsed overnight in buffer solution and transferred into osmium tetroxide for 12 hours to improve electron conductivity. Subsequently, segments were dehydrated in a graded 10% series of acetone solutions, for 10 minutes at each concentration between 10% to 80%, for 15 minutes at 90%; and two changes for 15 minutes in each of two changes of 100% acetone. The leaf segments were then dried in a critical point dryer. After drying, they were mounted on aluminium stubs, sputter coated with gold and examined with a JEOL JSM 820 scanning electron microscope at an accelerating voltage of 10 kV. Stomata on the lower and upper leaf surfaces of each of the 4 eucalypt species were photographed using 120 Ilford FP 4 film. The stomatal density was determined as the total number of stomates per mm2 for each of the 4 eucalypt species from 4 replicate photographs at x 250 magnification. The calibration bar in each photograph was used to calculate the field area. Preparation of solutions Two treatment solutions were used; 20% glycerol (v/v) as a preservative solution, and deionised water as a control. A biocide (10 mg/L available chlorine as dichloroisocyanurate) was added to each of the solutions. Ten replicates, each consisting of a cut stem in its own vase, were placed in a controlled environment at 24 ±1°C and 65 ±5% RH during experimentation using a completely randomised design. Initial and daily change in mass was recorded daily for 9 days. The cumulative uptake of solution (total amount of solution taken up), was calculated using the following formula: Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time in hours. A completely randomised design was used. Analysis of variance were conducted using the General Linear Model procedure with the statistical package Minitab, Version 11 (Anon. 1996). Mean separation was by the least significant difference (LSD) test at p=0.05.

Detailed Results The four eucalypt species, E. robusta, E. tetragona, E. bakeri and E. crucis, all had different morphological and anatomical characteristics (Tables 2.1 and 2.2).. E. robusta had fewer leaves per stem and no stomata on the upper surface of its leaves. In addition E. robusta had fewer larger vessels in its xylem. E. tetragaona and E. bakeri had thicker leaves than the other eucalypt species. Of the four eucalypt species that were studied, only E. tetragona showed well-defined layers of palisade mesophyll on both sides of the leaf (Plate 2.2), suggesting its adaptation to xerophytic

13

environments. This species also had a much thicker cuticle on both the upper and lower sides of its leaf than all other species. Table 2.1. Morphological characteristics of the leaves and stems of four Eucalyptus species. Means (± s.e.) are presented (20 replications except for leaf area where there were 10 replications). Species Leaf number

per 30 cm

stem

Leaf area

(mm2)

Total leaf area

per 30 cm stem

(mm2)

Leaf thickness

(mm)

Stem thickness

(mm)

E. robusta 6.60 ± 0.3 45.2 ± 2.8 294 ± 13 0.43 ± 0.01 5.32 ± 0.3

E. tetragona 11.8 ± 0.6 28.3 ± 1.3 333 ± 16 0.56 ± 0.01 7.97 ± 0.3

E. crucis 14.2 ± 0.7 21.3 ± 0.8 302 ± 14 0.44 ± 0.01 4.33 ± 0.1

E. bakeri 18.7 ± 1.6 5.51 ± 0.2 103 ± 9 0.55 ± 0.01 5.00 ± 0.1

Table 2.2. Anatomical characteristics of the leaves and stems of four Eucalyptus species. Means (± s.e.) are presented (6 replications except for stomatal density where there were 4 replications).

Stomatal density (number per mm2)

Cuticle layer thickness (µm)

Species

Lower Surface

Upper Surface

Lower Surfacer

Upper Surface

No. of 2ο xylem vessels

2ο xylem vessel size (µm)

E. robusta 282 ± 4

0

3.24 ± 0.21 3.53 ± 0.19 287 ± 2 59.0 ± 1.1

E. tetragona 186 ± 4 190 ± 7

11.4 ± 0.08 12.2 ± 0.16 481 ± 11 38.8 ± 2.0

E. crucis 200 ± 6 178 ± 6

3.15 ± 0.12 3.57 ± 0.17 417 ± 14 41.0 ± 1.7

E. bakeri 122 ± 7

129 ± 4 6.34 ± 0.24 6.67 ± 0.32 345 ± 7 37.3 ± 1.7

14

Plate 2.2. Part of a transverse section showing the leaf anatomy of Eucalyptus tetragona with palisade mesophyll on both sides of the leaf and a large oil gland on centre left of photograph. The four eucalypt species, E. robusta, E. tetragona, E. bakeri and E. crucis, were found to display differences in cumulative solution uptake of water but showed a lower and similar cumulative solution uptake of glycerol (Table 2.3). E. tetragona showed a consistently lower cumulative solution uptake of water than E. robusta and E. crucis every day for 9 days and a lower cumulative solution uptake than E. bakeri after Day 4. However, no strong correlation between solution uptake and either xylem area or the number of xylem vessels was observed. It is, however, possible that not all of the vessels observed in the transverse stem sections were involved in solution movement. Alternatively, other plant factors such as cuticle thickness and anatomical structure of the leaf may have contributed to the recorded differences in solution uptake behaviour among the four species. Table 2.3: Cumulative uptake of deionised water and 20% glycerol solution by four Eucalyptus species. Species Cumulative uptake of deionised water (gg fw-1) Day 1 Day 2 Day 3 Day 4 Day 5 Day 7 Day 9 E. robusta

0.68

1.59

2.27

3.13

4.00

5.63

6.70

E. tetragona

0.36

0.74

1.06

1.48

1.89

2.50

2.87

E. crucis

0.50

1.35

2.12

3.43

4.66

6.08

6.93

E. bakeri

0.27

0.50

0.98

1.89

2.80

5.31

7.35

Cumulative uptake of 20% glycerol solution (gg fw-1) E. robusta

0.67

0.88

0.95

0.97

1.00

1.03

1.06

E. tetragona

0.22

0.59

0.84

0.94

1.05

1.12

1.16

E. crucis

0.54

1.03

1.16

1.21

1.25

1.30

1.34

E. bakeri

0.27

0.92

1.07

1.10

1.14

1.18

1.23

LSD0.05

NS

0.096

0.162

0.199

0.248

0.353

0.433

15

Implications The uptake of preservation solutions requires fresh and healthy plant tissues. Stems that have been cut and poorly handled during transportation to the preservation shed are more likely to show variable or poor solution uptake. While different species adapted to xerophytic environments may have different morphological and anatomical characters, this may not necessarily affect the uptake of glycerol solution and hence preservation.

Recommendations Stems to be preserved by uptake should be handled in a similar way to those stems to be sold as fresh cut flowers. The time between harvest and preservation needs to be minimised. Stems should be recut before being placed into preservative solutions.

16

3. Do postharvest environmental factors affect the uptake of glycerol solution by Eucalyptus robusta? Introduction The postharvest conditions which plant material is subjected to during transportation, handling and processing all effect the uptake of preservation solutions such as glycerol. These factors also impact on the quality of the preserved material and the economics of processing. This chapter investigates the effect of various postharvest conditions on the rate of uptake, the amount of solution taken up and the relative change in mass of the plant material caused by solution uptake for cut stems of Eucalyptus robusta.

Objectives 1. Investigate the effect of prewilting during transportation and handling on the uptake of glycerol

solution by Eucalyptus robusta. 2. Investigate the effect of light intensity during processing on the uptake of glycerol solution by E.

robusta. 3. Investigate the effect of airspeed during processing on the uptake of glycerol solution by E.

robusta. 4. Investigate the effect of vapour pressure deficit during processing on the uptake of glycerol

solution by E. robusta. 5. Recommend to industry the environmental conditions which will enhance uptake of glycerol

solution by E. robusta.

Methodology General methodology for investigating the rate of uptake, the amount of solution taken up and the relative change in mass Stems of E. robusta foliage were harvested from a managed garden at The University of Queensland Gatton. In all experiments accept the prewilting experiment, stems were cut to 20 cm and held overnight at 5°C in buckets of deionised water covered with polyethylene sheets. For each of the experiments two treatment solutions were used; 20% glycerol (v/v) as a preservative solution, and deionised water as a control. A biocide of 10 mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Ten replicates, each consisting of a cut stem in its own vase, were placed in a controlled environment at 24 ±1°C and 65 ±5% RH during experimentation using a completely randomised design. Other environmental factors were imposed as detailed for each experiment. Solution uptake was recorded at 6, 12, 24, 30 and 48 hours and daily thereafter for up to 7 days unless otherwise stated. Observations were noted of leaf colour changes from green to brown as an indicator of glycerol accumulation. For each assessment the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up), and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0

17

Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours, and n-1 is the previous assessment time in hours. A factorial design was adapted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. Analyses for the treatment effects of prewilting were done 1, 3, 5 and 7 days after treatments were applied. Analyses for the treatment effects of the environmental factors used (light, vapour pressure deficit and airspeed) were done at 6, 24, 72 and 168 h after treatments were applied. A full set of statistical results pertaining to this chapter can be found in Appendix 4. Modifications for investigating the effect of prewilting during transportation and handling To simulate effects of transport and handling time prior to putting stems into preservative solutions cut E. robusta foliage stems were left out of the hydrating solution in the vase life room (20 ±2oC and 70 ±5% RH) for five prewilting times: • 0 hours (control). These stems were placed immediately into solution after weighing. • 6 hours, • 12 hours, • 24 hours and • 48 hours. Stems were cut in the field and wrapped in thin plastic film to prevent transpiration during transport to the laboratory. All stems were weighed immediately on arrival at the laboratory prior to prewilting treatments. On completion of prewilting treatments, stems were reweighed, and their water potential was determined using a pressure bomb described by Turner (1988). Stems were placed into the solutions and solution uptake was recorded daily for 7 days. After prewilting five replicates each were used for determining solution uptake rate, cumulative uptake and relative mass for deionised water and 20% glycerol solution treatments. Modifications for investigating the effect of light intensity during processing To investigate the effect of light intensity during processing, cut E. robusta foliage stems were subjected to four light regimes during solution uptake. The light regimes were: • normal room lighting from two 40 W Osram® white fluorescent lamps at 75 cm above the leaves

(10.5-15.0 µEinm-2 s-1, control), • continuous high light regime from four 36 W Philips® cool white fluorescent lamps at 25 cm

above the leaves (56-75 µEinm-2 s-1), • a 12 h on-off cycle with normal room lighting as described above, and • continuous darkness created by enclosing a section of the room with thick black polyethylene

sheets. Modifications for investigating the effect of airspeed during processing To investigate the effect of airspeed during processing cut E. robusta foliage stems were subjected to four different environments during solution uptake. The airspeed environments were: • undirected air movement, where stems were kept in an open environment (control),

18

• enclosed, where stems were surrounded by 60 cm high walls (0 ms-1), • directed low airspeed, where stems were placed 30 cm away from two fans (0.5 ms-1), and • directed high airspeed, where stems were placed 5 cm away from two fans (1.5 ms-1). Modifications for investigating the effect of vapour pressure deficit during processing To investigate the effect of vapour pressure deficit (VPD) during processing, cut E. robusta foliage stems were held in controlled rooms of different temperatures to create four different VPD environments during solution uptake. The VPD is the difference between the leaf and the atmospheric vapour pressure. The leaf vapour pressure at saturation and the atmospheric vapour pressure were recorded for each of the room conditions. The atmospheric vapour pressure in each room was obtained from hygrometric tables based on the difference between wet and dry bulb readings. The resulting environmental conditions for each of the temperature rooms are detailed in Table 3.1. Table 3.1. The vapour pressure deficit experienced by the cut stems of E. robusta foliage during uptake of solution in each of the temperature controlled rooms. Leaf vapour pressure was determined at saturation.

Tdry bulb (°C)

Twet bulb (°C)

Atmospheric vapour pressure

(mb)

Leaf vapour pressure

(mb)

Vapour pressure deficit (mb)

0.5 0.1 5.8 6.3 0.5 7.0 6.0 8.5 10.0 1.5

20.0 15.8 14.6 23.4 8.8 30.0 20.0 20.0 42.4 22.4

Detailed results Effect of prewilting simulating transportation and handling Stem mass was significantly reduced as the time that stems were left out of solution increased (Table 3.2). The reduction in mass approximately doubled with each doubling of the time stem were left out of solution. Stems left out of solution for 48 hours lost almost half of their initial weight (44.2%). The water potential of the stems was measured to determine the water stress to which they were subjected by the prewilting treatments. The control had the lowest water potential at –1717 MPa. The longer the stems were left out of solution the lower was their water potential. Table 3.2. The reduction in mass and water potential of cut E. robusta stems after different prewilting periods. Means followed by the same letter are not significantly different at p=0.05.

Prewilting time

Mass reduction (% initial weight)

Water potential (-MPa)

0 h (Control) 0.0 a 1717 a 6 h 4.1 b 2858 b

12 h 9.9 c 3350 c 24 h 21.8 d 4065 d 48 h 44.2 e 5565 e

LSD p=0.05 1.6 224 The type of uptake solution and the period of prewilting significantly influenced the uptake of solution. There was a strong interaction between these two factors. Initially stems that had been prewilted for 6 and 12 h took up solution most rapidly, although there was no significant difference between the control and the 6 h prewilt treatment (Table 3.3). After day 3 there was a significant difference in the uptake of solution by those stems that had been prewilted for 12 hours or less and those that had been prewilted for more than 24 hours.

19

Uptake of the glycerol solution by stems that had been prewilted for more than 24 hours was very poor throughout the experiment, being less than 0.9 gg-1 fw day-1 (Figure 3.1 (b). Consequently, their cumulative uptake was also low and their relative mass was significantly lower than that of the control stems and of those dehydrated for 6 and 12 hours before processing (Figures 3.2(a), 3.2(b), 3.3(a) and 3.3(b)). In contrast, stems that had been prewilted for 12 hours or less initially took up the glycerol solution relatively quickly at an average 0.75 gg-1 fw day-1 (Figure 3.1(b). However, the rate of uptake by these stems declined from day 3, and by day 4 their rate of glycerol uptake did not differ from those stems prewilted for a longer period. The cumulative uptake continued steadily for stems prewilted for 12 h or less in glycerol solution but leveled off at around 1.21 gg-1 fw after 5 days. In contrast, stems that had undergone the same treatments but were placed in the deionised water continued to take up solution consistently throughout the experiment. The relative mass of those stems that had had been prewilted for 12 hours or less increased initially or increased after an initial weight loss recorded on day 1 and then remained steady during the experiment. Whilst gain in relative mass was greater in those prewilted for 12 hours than in those prewilted for 6 hours, the difference was not significant. In these stems held in the glycerol solution, the resulting relative mass was in excess of 107% of the initial weight of the stems, indicating that the water in the stems had been replaced by the glycerol solution and had maintained their moisture. In contrast, stem that had been prewilted for 24 hours or more never regained their initial weight in either solution indicating that this pretreatment had caused irreparable damage to the plant tissues. Table 3.3: Effect over time of prewilting treatment on rate of solution uptake by stems of E. robusta. . Means followed by the same letter are not significantly different at p=0.05.

Prewilting Time

Day 1 Day 3 Day 5 Day 7

0 h (Control)

#0.5279 c 0.2241 b 0.1388 b 0.1476 b

6 h 0.5495 cd 0.2359 b 0.1794 b 0.1518 b

12 h 0.6268 d 0.2159 b 0.1599 b 0.1565 b

24 h 0.1529 b 0.0421 a 0.0240 a 0.0116 a

48 h 0.0342 a 0.0165 a 0.0152 a 0.0139 a

LSD p=0.05

0.0878 0.0463 0.0439 0.0214

# Values are in gg-1 fw d-1

20

Figure 3.1(a): Effect over time of prewilting treatment on rate of solution uptake by stems of E. robusta in deionised water (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h).

Figure 3.1(b): Effect over time of prewilting treatment on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h). Bars show LSD (p=0.05) between means for the interaction of prewilting treatment and solution for 4 selected observation times (1, 3, 5 and 7 days). No bar at a selected observation time indicates non-significance.

0

0.2

0.4

0.6

0.8

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Solu

tion

Upt

ake

Rat

e (g

g-1 fw

d-1

)

- Glycerol

0

0.2

0.4

0.6

0.8

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Sol

utio

n U

ptak

e R

ate

(gg-1

fw d

-1)

+ Glycerol

21

Figure 3.2(a): Effect of prewilting treatment on cumulative solution uptake by stems of E. robusta in deionised water. (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h).

Figure 3.2(b): Effect of prewilting treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution. (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h). Bars show LSD (p=0.05) between means for the interaction of prewilting treatment and solution for 4 selected observation times (1, 3, 5 and 7 days). No bar at a selected observation time indicates non-significance.

0

0.5

1

1.5

2

2.5

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Cum

ulat

ive

Solu

tion

Upt

ake

(gg-1

fw)

- Glycerol

0

0.5

1

1.5

2

2.5

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Cum

ulat

ive

Solu

tion

Upt

ake

(gg-1

fw)

+ Glycerol

22

Figure 3.3(a): Effect over time of prewilting treatment on relative mass of stems of E. robusta in deionised water (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h).

Figure 3.3(b): Effect over time of prewilting treatment on relative mass of stems of E. robusta in 20% glycerol solution (υ = control; ν = 6 h; σ = 12 h; λ = 24 h; x = 48 h). Bars show LSD (p=0.05) between means for the interaction of prewilting treatment and solution for 4 selected observation times (1, 3, 5 and 7 days). No bar at a selected observation time indicates non-significance. Effect of light intensity during processing Solution type and the light intensity regime to which the samples were subjected significantly affected the uptake of solution during the experiment. There was a strong interaction between these two factors (Figures 3.4(a) and 3.4(b)).

0

20

40

60

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There was no significant difference in the rate of uptake of either the deionised water or the glycerol solution initially at 6 hours. However, variation was observed in the uptake rates caused by the different light regimes. Uptake was most rapid under the high light, whilst in continuous darkness uptake was quite slow (Table 3.4). After 24 hours, the uptake rate of both solutions had decreased by about 60% based on the mean of all light intensities. This trend of declining uptake rate continued throughout the remainder of the experiment for cut stems E. robusta in the glycerol solution, and after 72 hours there was no difference in the uptake rate between the light intensity treatments. Uptake remained reasonably steady for the deionised water, despite the erratic uptake of the stems under the high light intensity. It is possible that the high light intensity environment may have affected the behaviour of the stomata. The cumulative uptake of solution was greatest under the high light regime, particularly for the deionised water (Figures 3.5(a) and 3.5(b)). At 24 hours the stems under the high light regime had accumulated 1.24 gg-1 fw or 31% more glycerol solution than the stems in normal light (control). After 72 hours there was no difference between the two treatments for the uptake of the glycerol solution. The stems subjected to the on-off cycle and to continual dark did not differ in their cumulative uptake of glycerol solution. Table 3.4: Effect over time of light treatment on rate of solution uptake by stems of E. robusta. Means followed by the same letter are not significantly different at p=0.05.

6 h 24 h 72 h 168 h

Control Normal R

#0.0708 c 0.0255 b 0.0124 a 0.0175 b

High Light Regime

0.0921 d 0.0289 b 0.0379 b 0.0179 b

12 h on-off cycle

0.0569 b 0.0201 a 0.0153 a 0.0141 b

Continuous dark

0.0264 a 0.0190 a 0.0116 a 0.0086 a

LSD p=0.05

0.0123 0.0045 0.0053 0.0040

# Values are in gg-1 fw d-1

24

Figure 3.4(a): Effect over time of light treatment on rate of solution uptake by stems of E. robusta in deionised water (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark).

Figure 3.4 (b): Effect over time of light treatment on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark). Bars show LSD (p=0.05) between means for the interaction between light treatment and solution for 4 selected observation times. No bar at a selected observation time indicates non-significance.

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Figure 3.5(a): Effect of light treatment on cumulative solution uptake by stems of E. robusta in deionised water (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark).

Figure 3.5(b): Effect of light treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark). Bars show LSD (p=0.05) between means for the interaction between light treatment and solution for 4 selected observation times (6, 24, 72and 168 h). No bar at a selected observation time indicates non-significance.

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Figure 3.6(a): Effect over time of light treatment on relative mass of stems of E. robusta in in deionised water (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark).

Figure 3.6(b): Effect over time of light treatment on relative mass of stems of E. robusta in 20% glycerol solution (υ = normal light intensity i.e. control; ν = high light intensity; σ = 12 hour cycle; λ = dark). Bars show LSD (p=0.05) between means for the interaction between light treatment and solution for 4 selected observation times (6, 24, 72and 168 h). No bar at a selected observation time indicates non-significance.

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Effect of airspeed during processing Solution type, airspeed and the interaction between these factors significantly influenced the solution uptake by cut stems of E. robusta. Irrespective of airspeed treatment, uptake of the solutions declined over time (Figures 3.7(a) and 3.7(b)). This was particularly noticeable for the glycerol solution and after 24 hours there was a significant difference in the uptake rate between the solutions. Stems in the low air speed environment consistently had the fastest solution uptake rate and those placed in the enclosed environment had a low uptake rate (Table 3.5). It is interesting to note that initially the uptake rate of stems in the high air speed environment was relatively fast, but showed a marked deceleration as the experiment progressed. After 24 hours the uptake rate of stems in this environment did not differ from either the low air speed environment or the control. Furthermore, after 72 hours, it did not differ from that in the enclosed environment. In considering the uptake rate of glycerol solution alone, a low airspeed resulted in the fastest uptake for the first 24 hours of the experiment, but after 72 hours the uptake rate did not differ from the control or the enclosed environment (Figure 3.7(b)). Cumulative uptake of glycerol solution was highest by stems of E. robusta experiencing a low airspeed (Figure 3.8(b)). Forty percentage of these stems turned greenish brown on the first day of the experiment, indicating that were accumulating glycerol in their leaves. By 72 hours, the leaves on all of the stems in this treatment had turned brown. By comparison stems in the control and high airspeed treatments only achieved this after 96 hours. Stems in the enclosed environment took more than 168 hours to show a change in leaf colour. As expected, there was no observed colour change in the leaves on stems in the deionised water. The change in relative mass was not significantly affected by the airspeed treatments (Figures 3.9 (a) and (b). Table 3.5: Effect over time of airspeed treatment on rate of solution uptake by stems of E. robusta. Means followed by the same letter are not significantly different at p=0.05.

6 h 24 h 72 h 168 h

Control #0.0553 b 0.0333 b 0.0237 b 0.0138 b

Enclosed 0.0349 a 0.0273 a 0.0127 a 0.0067 a

Low Air Speed

0.0701 d 0.0379 c 0.0284 c 0.0188 c

High Air Speed

0.0633 c 0.0353 bc 0.0120 a 0.0067 a

LSD P=0.05

0.0045 0.0040 0.0028 0.0020

# Values are in gg-1 fw d-1

28

Figure 3.7(a): Effect over time of airspeed treatment on rate of solution uptake by stems of E. robusta in deionised water (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1).

Figure 3.7(b): Effect over time of airspeed treatment on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1). Bars show LSD (p=0.05) between means for the interaction of airspeed treatment and solution at 4 selected times (6, 24, 72 and 168 h). No bar at a selected observation time indicates non-significance.

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Figure 3.8(a): Effect of airspeed treatment on cumulative solution uptake by stems of E. robusta in deionised water (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1).

Figure 3.8(b): Effect of airspeed treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1). Bars show LSD (p=0.05) between means for the interaction of airspeed treatment and solution at 4 selected times (6, 24, 72 and 168 h). No bars at a selected observation time indicates non-significance.

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Figure 3.9(a): Effect over time of airspeed treatment on relative mass of stems of E. robusta in deionised water (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1).

Figure 3.9(b): Effect over time of airspeed treatment on relative mass of stems of E. robusta in 20 % glycerol solution (υ = undirected control, 0.1 ms-1; ν = enclosed, 0 ms-1; σ = directed low airspeed, 0.5 ms-1; λ = directed high airspeed, 1.5 ms-1). Bars show LSD (p=0.05) between means for the interaction of airspeed treatment and solution at 4 selected times (6, 24, 72 and 168 h). No bars at the selected observation time indicate non-significance. Effect of vapour pressure deficit during processing Increasing the room temperature had the effect of increasing the vapour pressure difference between the leaf and the atmosphere (Table 3.1). The vapour pressure deficit had a significant effect on the

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solution uptake rate. At 72 hours and 168 hours the solution type and the interaction between solution type and the vapour pressure deficit also had a significant effect on solution uptake rate. As the vapour pressure deficit experienced by the cut stems of E. robusta increased, so too did the rate of solution uptake (Table 3.6). The uptake rate at 22.4 mb was significantly faster that at all other VPD levels for each of the recording times. Uptake rate was consistently less than 0.008 gg-1 fw h-1 for the 0.5 and 1.5 mb treatments for the entire experiment. Uptake rate of the glycerol solution was most rapid at 22.4 mb being as high as 0.066 gg-1 fw h-1 at the 6 hours recording (Figure 3.10(b)). However, by 72 hours there were no differences between the vapour pressure deficit treatments. Cumulative solution uptake was also higher for the larger vapour pressure deficits (Figures 3.11(a) and 3.11(b)). In fact, by 24 hours, 60% of the E. robusta stems at 2.4 mb were observed to have changed colour from green to brown, indicating that they had absorbed glycerol solution. In contrast the 8.8 mb treatment that had the greatest increase in relative mass due to the uptake of glycerol solution (Figures 3.12(a) and 3.12(b)). Table 3.6: Effect over time of vapour pressure deficit treatment on rate of solution uptake by stems of E. robusta, averaged over solutions. Means followed by the same letter are not significantly different at p=0.05.

6 h 24 h 72 h 168 h

22.4 mb #0.06539 c 0.04084 d 0.04095 c 0.01500 c

8.8 mb 0.04365 b 0.02190 c 0.01285 b 0.01027 b

1.5 mb 0.00625 a 0.00780 b 0.00670 a 0.00250 a

0.5 mb 0.00520 a 0.00244 a 0.00520 a 0.00240 a

LSD p=0.05

0.00998 0.00346 0.00528 0.00282

# Values are in gg-1 fw d-1

32

Figure 3.10(a): Effect over time of vapour pressure deficit treatment on rate of solution uptake by stems of E. robusta in deionised water (υ = 22.4 mb; ν = 8.8 mb; σ a= 1.5 mb; λ = 0.5 mb).

Figure 3.10(b): Effect over time of vapour pressure deficit treatment on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = 22.4 mb; ν = 8.8 mb; σ = 1.5 mb; λ = 0.5 mb). Bars show LSD (p=0.05) between means for the interaction of vapour pressure deficit treatment and solution for 4 selected observation times (6, 24, 72 and 168 h). No bar at a selected observation time indicates non-significance.

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Figure 3.11(a): Effect of vapour pressure deficit treatment on cumulative solution uptake by stems of E. robusta in deionised water (υ = 22.4 mb; ν = 8.8 mb; σ = 1.5 mb; λ = 0.5 mb)

Figure 3.11(b): Effect of vapour pressure deficit treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = 22.4 mb; ν = 8.8 mb; σ = 1.5 mb; λ = 0.5 mb). Bars show LSD (p=0.05) between means for the interaction of vapour pressure deficit treatment and solution for 4 selected observation times. (6, 24, 72 and 168 h). No bar at a selected observation time indicates non-significance

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Figure 3.12(a): Effect over time of vapour pressure deficit treatment on relative mass of stems of E. robusta in deionised water (υ = 22.4 mb; ν = 8.8 mb; σ = 1.5 mb; λ = 0.5 mb).

Figure 3.12(b): Effect over time of vapour pressure deficit treatment on relative mass of stems of E. robusta in 20% glycerol solution (υ = 22.4 mb; ν = 8.8 mb; σ = 1.5 mb; λ = 0.5 mb). Bars show LSD (p=0.05) between means for the interaction of vapour pressure deficit treatment and solution for 4 selected observation times (6, 24, 72 and 168 h). No bar at a selected observation time indicates non-significance.

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Discussion of results These experiments showed that the uptake of preservation solutions can be enhanced by modifying the environmental conditions prevailing during transportation, handling and processing. Uptake of solutions is via the transpiration stream. Therefore, factors that hasten transpiration are likely to improve the movement of solution through the stem to the leaves. These may possibly include both plant (e.g. prewilting) and environmental (e.g. light intensity, airspeed and vapour pressure deficit) factors. Preservation processing under controlled conditions can result in more rapid uptake and more uniform distribution of preservative solution through the plant material. This can be economical and produce a superior product. Prewilting After harvest, the cut stems of E. robusta continue to transpire and will dehydrate in the absence of a water supply, drawing the sap up the xylem vessels. Rehydration is then dependent on the establishment of a continuous water column in the xylem vessels between the leaf and the water supply. In an earlier study, Dubois and Joyce (1992) showed that prewilting jarrah foliage for 12 hours enhanced the initial rate of glycerol solution uptake. However, longer periods of prewilting resulted in an inability of the material to rehydrate and take up glycerol. In contrast this study indicated that there was no apparent benefit of prewilting of cut E. robusta stems on uptake of glycerol solution. There were, however, no apparent detrimental effects of prewilting for up to 12 hours. Prewilting for 24 hours or more in this study resulted in an inability of the material to rehydrate and take up glycerol, as shown by Dubois and Joyce (1992). It is likely that the solution was unable to sufficiently penetrate the xylem vessels and re-establish the water column. This problem may be overcome by recutting the stems, particularly if it is done under water, or by the addition of a wetting agent to the solution. However, given the degree of reduction in mass of the stems that had been prewilted for 24 and 48 hours (22.8 and 44.2% respectively), it is possible that the stem and leaf cells had been desiccated beyond recovery. Light intensity Light can affect stomatal activity in both a direct and an indirect manner. The stomata of most plants are open in response to light and closed in response to darkness. Thus, it can be expected that transpiration will be enhanced and consequently more solution taken up when more stomata are open. In this study, solution uptake increased with higher light intensity. Continuous lighting also improved solution uptake compared with the 12 hours on-off cycle. Airspeed The degree of airflow around a leaf influences the resistance of the boundary layer surrounding that leaf and ultimately its transpiration rate. The boundary layer is a region of reduced air movement adjacent to the leaf surface. As the airflow increases, the thickness of the boundary layer is reduced, as is its resistance, and the rate of transpiration is increased. The results showed that the low airspeed (0.5 ms-1) was enough to decrease the boundary layer resistance around the leaf and increase the uptake of solution compared with the control (undirected airflow) and a sheltered environment. The higher airspeed (1.5 ms-1) would have also had the effect of decreasing the boundary layer resistance, yet the total or cumulative solution uptake was reduced. It is likely that the reduction in boundary layer was so great that it caused desiccation and induced stomatal closure, thereby reducing transpiration. High airspeeds can also cause leaves to rub together resulting in mechanical damage to the leaf surfaces and stomata. Vapour pressure deficit The stomata of some species respond to changes in atmospheric humidity or vapour pressure deficit (VPD). The internal atmosphere of the leaf is generally at saturation while the external atmosphere is usually in an unsaturated condition. As the difference in vapour pressure between the internal and external atmospheres becomes greater to a reasonable limit, vapour will diffuse more rapidly through

36

the stomata. Consequently, solution would be taken up more rapidly. The vapour pressure deficit can be manipulated by altering the temperature and relative humidity of the environment. This experiment clearly demonstrated the influence of vapour pressure deficit on uptake of solution by E. robusta. Solution uptake rate was most rapid at 22.4 mb and the cumulative solution uptake was also high at a large vapour pressure difference. Thus, by increasing the atmospheric temperature and consequently reducing the relative humidity, the time required for the accumulation of the glycerol preserving solution is reduced.

Implications The uptake of preservative solutions can be enhanced by modifying the prevailing environmental conditions during transportation, handling and processing. This could result in more rapid and uniform solution uptake. Prewilting of cut stems of E. robusta foliage for up to 12 hours prior to processing does not affect the uptake of glycerol. However, prewilting offers no advantage over processing material immediately after harvesting. Cut stems of E. robusta foliage that are not processed within 24 hours will not take up sufficient preservative solution. Cut stems of E. robusta foliage can be encouraged to take up glycerol solution more rapidly by subjecting them to continuous high light intensities. Air circulation at low airspeeds can increase the uptake of glycerol solution by cut stems of E. robusta foliage. High airspeeds can be detrimental to the uptake of solution The uptake of glycerol solution by cut stems of E. robusta foliage is improved by increasing the vapour pressure difference between the leaf and the atmosphere. This can be achieved by increasing the temperature during processing. Uptake is very poor at or below 7°C. Uptake of glycerol needs to be completed within 3 days. Whilst most processors do not have elaborate environmentally controlled sheds in which to process cut stems of foliage like that of E. robusta, there are some very simple procedures and equipment which could be put in place to improve the economics and out-turn quality of uptake preservation.

Recommendations Stems of cut E. robusta foliage are best processed promptly and within 12 hours of harvest. High intensity lighting (e.g. 36 W Philips® cool white fluorescent lamps, to give 56-75 µEinm-2 s-1) should be installed approximately 25 cm above the foliage being processed and remain illuminated throughout the uptake process. Fans should be installed in the processing area and operated on low during processing.

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4. Is the uptake of preservative solution by Eucalyptus robusta and E. tetragona affected by the type of humectant used and its concentration? Introduction Glycerol is the humectant most commonly used for the preservation of cut flowers and foliages. It is liquid at room temperature and thus, when used in uptake preservation, maintains the plant material’s natural suppleness. Added to this is its ability to attract relatively large amount of water from the atmosphere. However, this can be a disadvantage at particularly high relative humidities, as high absorption of water can cause the preserved plant material to sweat. This in turn can encourage the growth of mould and thereby reduce the quality of the product. Therefore, in order to produce a good quality preserved product, it is essential to determine the most appropriate preservative solution, the optimum uptake and thus the optimum concentration. This chapter investigates the effect of humectant type and concentration on uptake of solution by Eucalyptus robusta and E. tetragona, with an emphasis on glycerol.

Objectives 1. Investigate the effect of glycerol concentration on uptake of solution by Eucalyptus robusta and

E. tetragona. 2. Investigate the effect of humectant type and concentration on uptake of solution by Eucalyptus

robusta and E. tetragona.

Methodology General methodology for investigating the rate of uptake, the amount of solution taken up and the relative change in mass Stems of Eucalyptus robusta foliage were harvested from a managed garden at The University of Queensland Gatton and returned to the laboratory in water. The stems of E. robusta were re-cut to 30 cm in length, each with 4 to 5 leaves and weighed. Foliage stems of E. tetragona were harvested from a commercial plantation known as “Brookvale Park” near Cecil Plains in the afternoon of the day prior to commencing the experiment. On return to the laboratory the stems of E. tetragona were allowed to stand in deionised water overnight, then re-cut under water to 25 cm in length with 8 to 10 leaves and weighed. Separate experiments were conducted for E. robusta and E. tetragona. For each species two separate experiments were conducted; one relating to the concentration of glycerol solution and the other investigating different humectants, as outlined below. Ten replicates per treatment, each consisting of a cut stem in its own vase, were placed in a controlled environment at 20 ±2°C and 50% RH and a light intensity of 60-70 µEinm-2s-1. A completely randomised design was used. Solution uptake and stem weight were recorded once daily over 7 days. For each assessment the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0

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Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time in hours. To calculate the actual amount of glycerol taken up, the total weight of solution taken up was multiplied by a factor based on the glycerol concentration and the specific gravity of glycerol (1.262). Thus for a 10% glycerol solution (v/v), the factor is 0.123, that is it is equivalent to a 12.3% glycerol solution (w/w). For a 20% glycerol solution (v/v), the factor is 0.240, and 0.351 for a 30% solution (v/v). A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results can be found in Appendix 16.5. Modifications for investigating the effect of glycerol concentration Three glycerol treatment solutions were used: 10%, 20% and 30% glycerol (with concentrations determined on a volume to volume basis), with deionised water used as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Modifications for investigating the effect of humectant type and concentration Three humectant treatment solutions were used; 20% glycerol, 10% glycerol plus 10% polyethylene glycol (PEG) and 20% PEG. All concentrations were on a volume to volume basis. Deionised water was used as a control treatment. A biocide of 15mg/L available chlorine as dichloroisocyanurate was added to each of the solutions.

Detailed results Effect of glycerol concentration Initially there was no significant difference in uptake rate between the various treatment solutions for cut stems of either E. robusta or E. tetragona (Figures 4.1 and 4.2). However, uptake was more than twice as rapid in E. robusta (average 0.013 gg-1 fw h-1) than E. tetragona (average 0.005 gg-1 fw h-1). The uptake rate in the cut stems peaked on Day 2 for all of the glycerol treatments for E. robusta and for the 30% glycerol solution treatment for E. tetragona. The uptake rate in E. tetragona for the other glycerol treatments peaked at Day 3. In contrast the uptake of the deionised water was more constant throughout the experiment. The uptake rate of the 20% and 30%glycerol solutions by E. robusta dropped off fairly quickly after peaking at Day 2. This limited the cumulative uptake of these solutions from Day 4 and Day 3 respectively (Figure 4.3). A trend emerged on Day 2 in the accumulation of the glycerol solutions by E. robusta in relation to the solution’s concentration. The stronger the solution, the lower the uptake that occurred. From Day 3, there was a significant difference in the amount of glycerol solution accumulated between the three concentrations for this species. However when the actual amount of glycerol accumulated by stems of E. robusta is considered, in the initial stages of the experiment, this trend was reversed (Figure 4.5 and Table 4.1). That is, on Days 1 and 2, more glycerol was taken up when the 30% solution was used, than for the 20% solution, and the

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least for the 10% solution. Throughout the experiment the least glycerol was accumulated from using the 10% solution, but on Day 3 the situation with the 20% and 30% solutions switched. Whilst there was no significant difference between these two treatments at this time, by Day 4 significantly more glycerol was accumulated by E. robusta stems in the 20% solution than in the 30% solution. Table 4.1. Cumulative uptake of glycerol compared with total solution uptake by for E. robusta (gg-1

fw). Different letters within each column show significant differences in uptake for either total solutions or glycerol (LSD p=0.05). Glycerol solution Uptake of: Day 1 Day 2 Day 4 Day 6

10% Solution 0.300 1.085 B 2.182 C 2.561 C

Glycerol 0.037 a 0.133 a 0.268 a 0.329 a

20% Solution 0.295 1.003 B 1.553 B 1.650 B

Glycerol 0.071 a 0.241 b 0.373 c 0.396 c

30% Solution 0.347 0.757 A 0.948 A 1.0286 A

Glycerol 0.122 b 0.266 b 0.333 b 0.361 b The observed cumulative uptake of the glycerol solutions and glycerol per se, was similar for cut stems of E. tetragona (Figure 4.4). However, ultimately less solution was accumulated when compared to E. robusta. Overall there was a steady accumulation of all treatment solutions, with no difference occurring between treatments until Day 5. From Day 5, the cumulative uptake of the 30% glycerol solution by stems of E. tetragona was significantly less than for all other treatment solutions, this trend already being evident from Day 3. The same trend existed in relation to cumulative uptake and solution concentration as for E. robusta. There was no significant difference between the 10% and 20% solutions in the amount of solution accumulated during the experiment, although slightly more of the 10% solution was taken up. When the actual uptake of glycerol by E. tetragona was calculated, significantly less glycerol was accumulated when a 10% solution was used as the uptake treatment (Figures 4.6 and Table 4.2). Despite the significantly lower accumulation of the 30% glycerol solution, there was no significant difference between the 20% and 30% solutions in terms of actual glycerol accumulated from Day 3. Table 4.2. Cumulative uptake of glycerol compared with total solution uptake by for E. tetragona (gg-1 fw). Different letters within each column show significant differences in uptake for either total solutions or glycerol (LSD p=0.05). Glycerol solution Uptake of: Day 1 Day 2 Day 4 Day 6

10% Solution 0.140 0.316 0.816 1.086 B

Glycerol 0.017 a 0.040 a 0.100 a 0.134 a

20% Solution 0.114 0.311 0.816 1.004 B

Glycerol 0.027 a 0.075 b 0.196 b 0.241 b

30% Solution 0.133 0.334 0.550 0.640 A

Glycerol 0.047 b 0.117 c 0.193 b 0.225 b With the exception of the deionised water treatment, the relative mass recordings for E. robusta were relatively high (>100%) and showed a general increase during the experimental period (Figure 4.7). The relative mass in all of the glycerol treatments and the deionised water control dropped on Day 6,

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with no significant difference between treatments. This possibly reflects a change in prevailing environmental conditions at that time as the stems in the glycerol solutions recovered by Day 7. In contrast, the relative mass of E. tetragona stems declined during the first half of the experiment irrespective of treatment solution (Figure 4.8). Stem in deionised water continued this trend for the remainder of the experiment, whereas stems in the glycerol treatments showed signs of steadying around the 85% mark by Day 7. Stems treated with the 30% glycerol solution had reached this level at Day 3, while for those in the 10% and 20% solutions this occurred around Day 6.

Figure 4.1: Effect over time of glycerol concentration on rate of solution uptake by foliage stems of E. robusta (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.2: Effect over time of glycerol concentration on rate of solution uptake by foliage stems of E. tetragona (υ = deionised water i.e. control: ν = 10% glycerol ; σ = 20%glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

Figure 4.3: Effect of glycerol concentration on cumulative solution uptake by foliage stems of E. robusta (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.4: Effect of glycerol concentration on cumulative solution uptake by foliage stems of E. tetragona(υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.5: Effect of solution concentration of glycerol on the cumulative uptake of glycerol by foliage stems of E. robusta (ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

Figure 4.6: Effect of solution concentration of glycerol on the cumulative uptake of glycerol by foliage stems of E. robusta (ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.7: Effect of glycerol concentration on relative mass of foliage stems of E. robusta (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

Figure 4.8: Effect of glycerol concentration on relative mass of foliage stems of E. tetragona (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Effect of humectant type and concentration This experiment showed that the uptake rate of deionised water by each of the two eucalypt species was relatively constant during the experiment (Figures 4.9 and 4.10). The average uptake rate was approximately 44% faster for E. robusta (0.021 gg-1 fw h-1) than for E. tetragona (0.014 gg-1 fw h-1). It is interesting to note that for E. robusta, the initial rate of solution uptake was faster for stems in the humectant treatments than in the deionised water control (Figure 4.9). However the rate of uptake of the humectants had slowed dramatically by Day 3, at which time there was no significant difference between the humectant treatments and the average uptake rate was 0.006 gg-1 fw h-1. This trend continued throughout the remainder of the experiment such that at Day 7 the uptake rate had slowed to 0.002 gg-1 fw h-1. In contrast, the uptake rate of humectant solutions by E. tetragona peaked at Day 2 (average 0.013 gg-1 fw h-1) before reducing to 0.003 gg-1 fw h-1 on Day 7 (Figure 4.10). There was little variation in uptake rate between the different humectant treatment solutions. Given the consistent rates of deionised water uptake throughout the experiment, both E. robusta and E. tetragona had a steady increase in recorded cumulative uptake of deionised water (Figures 4.11 and 4.12). By Day 7, E. robusta had accumulated 3.402 gg-1 fw and E. tetragona, 2.382 gg-1 fw. E. robusta stems, however, took up significantly less of the humectant solutions than of the deionised water (Figure 4.11). On Days 2 and 3, stems of E. robusta had taken up more of the solutions containing glycerol than that of PEG alone. Whilst this trend continued, from Day 4 there was no significant difference between the three humectant treatments in the amount of solution accumulated by E. robusta stems. By Day 7 this had reached an average of 1.478 gg-1 fw, or 44% of the amount taken up by stems in the control treatment. A similar trend of reduced uptake of humectant treatments as compared to the control was observed for E. tetragona (Figure 4.12). However, in contrast to E. robusta, the amount of 20% glycerol solution taken up by the stems of E. tetragona (Day 7, 1.422 gg-1 fw) was 47% more than that of the humectant solutions containing PEG (Day 7, average 0.970 gg-1 fw). There was no significant difference in cumulative uptake of PEG alone or in combination with glycerol, at the concentrations used. The relative mass of E. robusta stems in deionised water was very steady throughout the experiment and averaged 100.9% (Figure 4.13). At Day 1, there was no difference in the relative mass of stems treated with the solutions 20% PEG or the combination of 10% PEG and 10% glycerol. However, the relative mass of those stems in 20% PEG solution dropped dramatically to a low at Day 4 of 83.5%. These stems then recovered to a relative mass of 86.2% by Day 7. In contrast stems treated with solutions containing glycerol showed a gradual increase in relative mass during the experimental period, with no significant differences between treatments from Day 3 (average 97.5%) to Day 7 (average 101.6%). The control stems of E. tetragona also had a steady relative mass of 100.7% for the first 6 days but then dropped to 98.0% on Day 7 (Figure 4.14). At Day 1, the humectant treated stems had relative masses in the range of 94.8 to 96.4%, with the 20% PEG treated stems being significantly lower than the other treatments. All of the humectant treatments then recorded a reduction in relative mass to Day 3. This was particularly marked in the stems in the 20% PEG solution and the 10% PEG plus 10% glycerol solution. These stems had a relative mass of 77.5% (average) at Day 3 and remained around this level for the rest of the experiment. There was, however, a slight rise in the relative mass of the 10% PEG plus 10% glycerol solution treated stems. The stems in the 20% glycerol solution recorded their lowest relative mass on Day 3 (92.4%) and then recovered to an average of 94.5% during Days 4 to 7.

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By the completion of the experiment there was distinct and uniform discoloration of the leaves on stems of E. robusta in the 20% glycerol solution, yet they had maintained their flexibility. Leaves subjected to the 20% PEG solution showed similar discoloration but with greater intensity around the veins and they had become dry and brittle. The combination treatment resulted in intermediate discoloration between the two solutions used separately and flexibility had been maintained but not to the same degree as in the leaves treated with glycerol alone. Treatment with deionised water did not affect the flexibility of the leaves or cause discoloration after 7 days.

Figure 4.9: Effect over time of humectant treatment on rate of solution uptake by foliage stems of E. robusta (υ = deionised water i.e. control; ν = 20% glycerol; σ = 20% PEG; λ =10% glycerol + 10% PEG). Bars show LSD (p=0.05) among means. No bar at an obsevation time indicates non-

significance. Figure 4.10: Effect over time of humectant treatment on rate of solution uptake by foliage stems of E. tetragona (υ = deionised water i.e. control; ν = 20% glycerol; σ = 20% PEG; λ =10% glycerol + 10%

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PEG). Bars show LSD (p=0.05) among means. No bar at an obsevation time indicates non-significance.

Figure 4.11: Effect over time of humectant treatment on cumulative solution uptake by foliage stems of E. robusta (υ = deionised water i.e. control; ν = 20% glycerol; σ = 20% PEG; λ = 10% glycerol + 10% PEG). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.12: Effect over time of humectant treatment on cumulative solution uptake by foliage stems of E. tetragona (υ = deionised water i.e. control; ν = 20% glycerol; σ = 20% PEG; λ = 10% glycerol+ 10% PEG). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance

Figure 4.13: Effect over time of humectant treatment on relative mass of foliage stems of E robusta (υ = deionised water i.e. control; ν = 20% glycerol; σ = 20% PEG; λ = 10% glycerol+ 10% PEG). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance.

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Figure 4.14: Effect over time of humectant treatment on relative mass of foliage stems of E. tetragona (υ = deionised water i.e. control; ν = 20% glycerol; σ =20% PEG; λ = 10% glycerol + 10% PEG). Bars show LSD (p=0.05) among means. No bar at an observation time indicates non-significance. Discussion of results Effect of glycerol concentration Whilst there were very similar trends in the uptake of the glycerol solutions between the two species of eucalypts, the actual figures are quite different. E. robusta took up more than twice as much solution than E. tetragona, and did so much faster. While the uptake rate peaked at Day 2 for E. robusta, compared with at Day 3 for E. tetragona, uptake tended to cease around days 3 to 4. In contrast, uptake in stems of E. tetragona continued, in most cases, throughout the experiment. This reflects the anatomical differences noted in Chapter 3. Effect of humectant type and concentration It is interesting to note that in this experiment, the amount of the solutions accumulated by each of the species is quite similar but the pattern of rate of uptake was different. The trend in uptake rate in E. robusta also differed from that observed in the glycerol concentration experiment. In the glycerol concentration experiment uptake rates were initially low and then peaked at Day 2, whereas in this experiment uptake rates were high on Day 1 and then dropped. Similar peaking around Day 2-3 was observed for E. tetragona in both experiments. The results would suggest that the PEG molecule is too large to be satisfactorily taken up by E. tetragona to have any preservative effect (as noted by the significant drop in relative mass). Similarly PEG did not provide any benefits in E. robusta over a straight glycerol solution in terms of maintaining relative mass and indeed it had a detrimental effect on the quality of the foliage.

Implications Glycerol is an appropriate preservative solution for E. robusta and E. tetragona. The most rapid accumulation of glycerol in stems of E. robusta and E. tetragona is achieved with a 20% glycerol solution. Uptake of glycerol solutions by E. robusta is reduced after 4 days. PEG, at the concentrations and in the combination with glycerol tested, is an inappropriate humectant for the preservation of E. robusta and E. tetragona, as it caused discoloration and loss of flexibility of the foliage.

Recommendations E. robusta and E. tetragona should be preserved using a 20% glycerol solution. Uptake should be completed within 4 days. Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of E. robusta and E. tetragona.

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5. Do adjuvants affect the uptake of glycerol solution by Eucalyptus robusta and E. tetragona? Introduction In the context of preserving cut flowers and foliages, the term ‘adjuvants’ applies to chemicals that, when added to the preserving solution, improve the uptake or penetration of that solution into the plant material. Wetting agents or surfactants are a special type of adjuvant used to lower the surface tension of a liquid, thus increasing its spreading or wetting properties. In the case of uptake preservation, this can result in accelerated uptake of the solution and improved preservation of the plant material. This has obvious benefits to the industry. However, depending on the effective electrical charge on the molecules of adjuvant, they can interact with charged sites in the plant tissue, affecting its function. Anionic adjuvants have a negative charge and may react with the positively charged sites in the plant tissue. On the other hand non-ionic adjuvants do not have a charge, and therefore, will not interact with either the positively or negatively charged sites in plant tissue. Likewise non-ionic adjuvants are less reactive with other constituents of solutions. This chapter investigates the potential effectiveness of adjuvants on improving the uptake of glycerol solution by cut stems of Eucalyptus robusta and E. tetragona.

Objectives 1. Investigate the effectiveness of wetting agents on improving the uptake of glycerol solution by

Eucalyptus robusta and E. tetragona.

Methodology The effectiveness of wetting agents in improving the uptake of glycerol solution by E. robusta and E. tetragona was investigated using Agrimul PG 2067 and Geropon SDS (dioctylsulphosuccinate). These two commercial wetting agents were added at 100 mg mL-1 to the uptake solutions of 20% glycerol (v/v) as a preservative solution and deionised water used as a control, prior to commencement of the experiment. A biocide of 10 mg/L available chlorine as dichloroisocyanurate was also added to each of the solutions. Agrimul PG 2067 is a non-ionic surfactant derived from glucose, while Geropon SDS is an anionic surfactant. Stems of E. robusta foliage were harvested from a managed garden at The University of Queensland Gatton. On returning harvested plant material to the laboratory, stems were re-cut to 30 cm under water and placed in treatment solutions. Foliage stems of E. tetragona were harvested from a commercial plantation known as “Brookvale Park” near Cecil Plains in the afternoon of the day prior to commencing the experiment. On return to the laboratory, each stem was re-cut under water to 30 cm in length and allow to stand in deionised water overnight. Ten replicates, each consisting of a cut stem in its own vase, were placed in a vase life room at 20 ±2°C and 65 ±5% RH during experimentation. A completely randomised design was used. Solution uptake was recorded once daily over 7 days. The pH of the solution was measured twice at the start of the experiment and a mean pH calculated (Table 5.1).

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Table 5.1 Mean pH of solutions used in wetting agent experiments. pH Deionised (DI) water 4.6 DI + Agrimul PG 2067 5.1 DI + Geropon SDS 5.1 Glycerol solution 4.2 Glycerol solution + Agrimul PG 2067 4.0 Glycerol solution + Geropon SDS 4.7 For each of the five assessment times (days 1,2,3,5 and 7) the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours, and n-1 is the previous assessment time in hours. A factorial design was adapted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. Analyses of treatment effects were done at all observation times. A full set of statistical results pertaining to this chapter can be found in Appendix 6.

Detailed results This experiment showed that the solution type, the addition of an adjuvant to the solution and the plant species all differentially influence the uptake of solution. There was a significant interactive effect between the solution type and the adjuvant added. This effect tended to show as a reduced uptake of deionised water in the presence of an adjuvant, whilst the adjuvants did not alter the uptake of the glycerol solution. There was also a significant interaction between the solution type and the species of eucalypt. Overall, the solution uptake rate was fastest for the control stems, followed by those treated with Agrimul and Geropon SDS, respectively. The differences between the treatments, however, were not always significant (Table 5.2). The solution uptake rates for all adjuvant treatments for both E. robusta and E. tetragona in glycerol solution peaked at Day 2 and had become very slow by Day 5 (Figures 5.1(b) and 5.1(d)). Initially, the uptake rate and cumulative uptake by E. robusta was twice that observed for E. tetragona (Tables

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5.3 and 5.4). However, on Day 3 there was no significant difference between the species in rate of solution uptake. Cut stems of E. robusta took up significantly more solution than stems of E. tetragona, the mean cumulative solution uptake by Day 7 being 1.57 and 0.95 gg-1fw respectively. For each of the species, the stems in the control solutions of deionised water took up more solution than those stems in the treatment solutions containing the adjuvants (Figures 5.2(a) and5.2(c)). Those stems subjected to a solution with Geropon SDS added, accumulated the least amount of solution. Stems of both species held in solutions containing Geropon SDS had the greatest reduction in relative mass (Figures 5.3(a), 5.3(b), 5.3(c) and 5.3(d)). These stems were severely effected by the presence of this adjuvant and rapidly wilted. They later displayed signs of necrosis and became very brittle. The relative mass of E. tetragona stems held in the control solutions and solutions with Agrimul also declined. In contrast, cut stems of E. robusta held in these solutions maintained a fairly constant relative mass, which was close to 100% up to day 5. Table 5.2: Means for the effect over time of adjuvants on solution uptake rate by stems of Eucalyptus species, averaged over the solutions. Means followed by the same letter are not significantly different at p=0.05.

Adjuvant Day 1 Day 2 Day 3 Day 5 Day 7

Control #0.2561 b 0.3559 0.2326 b 0.1636 c 0.1330 b

Agrimul 0.2535 b 0.3224 0.1956 a 0.0909 b 0.0534 a

Geropon SDS

0.2064 a 0.3158 0.1803 a 0.0432 a 0.0493 a

LSD p=0.05

0.0430 NS 0.0367 0.0221 0.0160

# gg-1 fw d-1 Table 5.3: Means for the effect over time of species on rate of solution uptake, averaged over adjuvants and solutions. Means followed by the same letter are not significantly different at p=0.05.

Day 1 Day 2 Day 3 Day 5 Day 7

E. robusta #0.3192 b 0.4421 b 0.1991 0.1291 b 0.1115 b

E. tetragona 0.1581 a 0.2206 a 0.2065 0.0693 a 0.0456 a

LSD P=0.05

0.0351 0.0357 NS 0.0181 0.0130

# gg-1 fw d-1

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Table 5.4: Means for the effect of species on cumulative solution uptake, averaged over adjuvants and solutions. Means followed by the same letter are not significantly different at p=0.05.

Day 1 Day 2 Day 3 Day 5 Day 7

E. robusta #0.3192 b 0.7613 b 0.9604 b 1.2401 b 1.5666 b

E. tetragona 0.1581 a 0.3787 a 0.5852 a 0.8066 a 0.9548 a

LSD p=0.05

0.0351 0.0494 0.0530 0.0683 0.0838

# gg-1 fw

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Figure 5.1(a): Effect over time of adjuvants on rate of solution uptake by stems of E. robusta in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.1(b): Effect over time of adjuvants on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for the interaction of adjuvant, species and solution for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

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Figure 5.1(c): Effect over time of adjuvants on rate of solution uptake by stems of E. tetragona in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.1(d): Effect over time of adjuvants on rate of solution uptake by stems of E. tetragona in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for the interaction of adjuvant, species and solution for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

0

0.1

0.2

0.3

0.4

0.5

0.6

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Solu

tion

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ake

Rat

e (g

g-1 fw

d-1

)

- Glycerol

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0.1

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d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Solu

tion

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ake

Rat

e (g

g-1 fw

d-1

)

+ Glycerol

56

Figure 5.2(a): Effect of adjuvants on cumulative solution uptake by stems of E. robusta in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.2(b): Effect of adjuvants on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for the interaction of adjuvant, species and solution. for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

0

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1

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d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

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ake

(gg-1

fw)

- Glycerol

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d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

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ulat

ive

Solu

tion

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ake

(gg-1

fw)

+ Glycerol

57

Figure 5.2(c): Effect of adjuvants on cumulative solution uptake by stems of E. tetragona in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.2(d): Effect of adjuvants on cumulative solution uptake by stems of E. tetragona in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for significant interactions of adjuvant, species and solution for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

0

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(gg-1

fw)

- Glycerol

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3

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Cum

ulat

ive

Solu

tion

Upt

ake

(gg-1

fw )

+ Glycerol

58

Figure 5.3(a): Effect over time of adjuvants on relative mass of stems of E. robusta in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.3(b): Effect over time of adjuvants on relative mass of stems of E. robusta in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for significant interactions of adjuvant, species and solution for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

60

70

80

90

100

110

120

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Rel

ativ

e M

ass

(%)

- Glycerol

60

70

80

90

100

110

120

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Rel

ativ

e M

ass

(%)

+ Glycerol

59

Figure 5.3(c): Effect over time of adjuvants on relative mass of stems of E. tetragona in deionised water (υ = deionised water without adjuvant; ν = Agrimul; σ = Gerapon SDS).

Figure 5.3(d): Effect over time of adjuvants on relative mass of stems of E. tetragona in 20% glycerol solution (υ = 20% glycerol without adjuvant; ν = Agrimul; σ = Gerapon SDS). Bars show LSD (p=0.05) between means for significant interactions of adjuvant, species and solution for 5 selected observation times (1,2,3,5 and 7 days). No bar at a selected observation time indicates non-significance.

60

70

80

90

100

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120

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

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ass

(%)

- Glycerol

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d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

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(%)

+ Glycerol

60

Discussion of results The two commercial wetting agents tested in this study, Agrimul PG 2067 and Geropon SDS, did not improve the uptake of the glycerol solution by cut stems of either E. robusta or E. tetragona. In fact, the addition of Geropon SDS to the uptake solutions caused rapid decline in the quality of the stems with the foliage wilting and becoming brittle within a few days due to cell death. It is possible that, given that the adjuvants or wetting agents used, were both lipophilic and hydrophilic, they may have disturbed the structure of the lipid-based membranes in the plant cells thereby resulting in the decline observed. It is also possible that the anionic adjuvant, Agrimul PG 2067, may have also chelated catioinc ions or compounds of metabolic importance. It is interesting to note that both of these adjuvants had been effective in improving the uptake of glycerol solution by eucalypt foliage when used for immersion preservation. However it was observed in an earlier study that Agral 600 at 0.015% and 0.03% also did not increase glycerol solution uptake by E. cinerea (Campbell, 1996).

Implications The commercial adjuvants, Agrimul PG 2067 and Geropon SDS, at the concentrations tested, should not be added to a glycerol solution during uptake preservation of cut stems of either E. robusta or E. tetragona. The addition of Agrimul at this concentration does not provide any benefits and the addition of Geropon SDS was detrimental to the stems. The uptake of the glycerol solution varies between different species of Eucalyptus. It appears that preservation of cut stems of E. robusta by the uptake process will be completed more rapidly than by E. tetragona stems.

Recommendations Further experimentation is required to determine whether the use of adjuvants can provide any benefit to the uptake preservation process for E. robusta and E. tetragona. Such experimentation should consider the use of Agrimul PG 2067 and Geropon SDS at lower concentrations, as well as other adjuvants, like Agral 600. Testing needs to be done on a species by species basis.

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6. Can fluorescent dyes be used to characterise the uptake of glycerol solution by Eucalyptus robusta? Introduction Dyes can be added to uptake solutions to assist in characterising the movement of the uptake solution through the plant material. The uptake solution enters the plant material at the base of the stem through large vessels. The solution moves through the transpiration stream and into the leaves. Once in the leaves, the solution is able to move into small spaces between the cells, known as the apoplast, and to other regions of the leaf. The movement of solution through the apoplast can be investigated using the yellowish-green dye pyranine (1, 3, 6, 8-pyrenetetrasulphuric acid). Pyranine is highly soluble in water and has low affinity for cell wall components. Strugger (1974 cited by Canny, 1995) demonstrated that the dye spreading to the leaf surface in anticlinal walls was visible under a fluorescent microscope. This chapter characterises the movement of glycerol solution through cut stems of Eucalyptus robusta by using a fluorescent dye.

Objectives 1. Investigate the effect of dye on the uptake of glycerol solution by Eucalyptus robusta. 2. Characterise the movement of glycerol solution through cut stems of E. robusta.

Methodology General methodology for investigating the rate of uptake, the amount of solution taken up and the relative change in mass Stems of E. robusta foliage were harvested from a managed garden at The University of Queensland Gatton College. Stems were cut to 30 cm and held overnight at 5°C in buckets of deionised water covered with polyethylene sheets. For each assessment the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time and n is the assessment time in hours and n-1 is the previous assessment time in hours.

62

A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results pertaining to this experiment can be found in Appendix 7. Modifications for investigating the effect of dye on the uptake of glycerol solution Stems were placed in one or other of the two treatment solutions; 20% glycerol (v/v) as a preservative solution and deionised water as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. The fluorescent dye, pyranine (0.1% v/v) had been added to half of the vases of each of the solutions. Three replicates, each consisting of a cut stem in its own vase, were placed in the two experimental light regimes: • normal room lighting from two 40 W Osram® white fluorescent lamps at 75 cm above the leaves

(10.5-15.0 µEinm-2 s-1, control), or • continual high light regime from four 36 W Philips® cool white fluorescent lamps at 25 cm above

the leaves (56-75 µEinm-2 s-1), in a controlled environment at 20.5°C and 65% RH during experimentation. A completely randomised design was used. Solution and stem weights were recorded at 0 and 24 hours and the solution uptake determined. The degree of fluorescence in the leaves was observed after 24 h using an epifluorescence microscope fitted with a H2 filter tube and using UV light. The degree of fluorescence was scored 1 for no dye, 2 for slight fluorescence, 3 for moderate fluorescence, 4 for more than moderate fluorescence and 5 for very intense fluorescence. Modifications for characterising the movement of glycerol solution Stems were placed in one of the two treatment solutions; 20% glycerol (v/v) as a preservative solution and deionised water as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Four fluorescent dye treatments, using pyranine (0.1% v/v), were imposed on 10 replicates of each of the above solution treatments. The dye treatment was added to one set of the solution treatment vases at the time of establishing the experiment (Day 0), to another set after 1 day and to another after 2 days. No dye was added to the fourth set of vases. All vases were kept in a controlled environment at 20.5°C and 65% RH during experimentation. A completely randomised design was used. Solution and stem weight were recorded at the time of establishment and for each 24 hour period up to and after the dye was added. In the case of no dye being added, these measurements were made every 24 hours for a total of 3 days. The solution uptake and relative mass were determined. The degree of fluorescence in stem sections and on 5 leaves (including the leaf blade and the mid rib) was observed 24 hours after adding the dye using an epifluorescence microscope fitted with a H2 filter tube and using UV light. The degree of fluorescence was scored 1 for no dye, 2 for slight fluorescence, 3 for moderate fluorescence, 4 for more than moderate fluorescence and 5 for very intense fluorescence. The score for the mid ribs was averaged across the 5 leaves.

Detailed results Effect of dye on the uptake of glycerol solution The presence of the fluorescent dye, pyranine, did not affect the uptake of solution as indicated by the uptake rate and the relative mass of the stems after 24 h. As expected over such a short timeframe of only 24 h, the solution type had no effect on uptake rate although it did effect the relative mass of the stems (Tables 6.1 and 6.2). Stems in the glycerol solution were significantly heavier because of the greater specific gravity of glycerol (relative mass:

63

102.41%) than those stems in deionised water (relative mass: 101.1%), even though they had both taken up the same amount of solution (0.80 gg-1fw for the 24-hour period). Light intensity had a significant effect, with the uptake of solution being faster under the high light regime (1.00 gg-1fw d-1) than under normal room light. It should be noted, however, that the interaction between light intensity and solution type was significant: high light significantly increased the rate of uptake of deionised water compared with that of normal light but had no significant effect on the rate of uptake of 20% glycerol solution (Table 6.3). The presence of 0.1% pyranine in the solutions had no significant effect on either the rate of solution uptake or on relative mass (Tables 6.1 and 6.2). The interaction of dye treatment and solution type on rate of solution uptake was also not significant (Appendix 7, Table 16.7.2). However, there was a significant interaction between dye treatment and light intensity for relative mass: in the presence of dye, relative mass increased significantly under high light, whereas in the absence of dye the difference between high and normal light treatments was not significant (Table 6.4). In terms of the fluorescence score, both the presence of dye and the solution type treatments had a significant effect (Table 6.5). There was also a significant interaction between these two factors (Appendix 7, Table 16.7.7). In the absence of dye, all stems received a score of 1. That is, there was no detectable dye in the leaves. In contrast, the average score of those stems exposed to pyranine was 3.4, indicating that the leaves showed a slightly higher than moderate degree of fluorescence when observed under the fluorescence microscope (Table 6.5). It is interesting to note that even though there was no significant difference in the amount of deioinsed water or 20% glycerol solution taken up by the stems, those stems in the glycerol solution had a significantly higher fluorescence score. Table 6.1: Means for the effects of dye treatment, light intensity and solution type on the rate of solution uptake by foliage stems of E. robusta.

Dye Treatment

Solution Uptake

Rate

Light Intensity Solution Uptake

Rate

Solution Solution Uptake

Rate Dye #0.80 High Light 1.00 a 20% Glycerol 0.79

No Dye 0.79 Normal Light 0.59 b DeionisedWater 0.80 LSD

(p=0.05)

NS

0.21

NS # Values are gg-1 fw d-1 Table 6.2: Means for the effects of dye treatment, light intensity and solution type on relative mass of foliage stems of E. robusta

Dye Treatment

Relative Mass

Light Intensity Relative Mass

Solution Relative Mass

Dye #101.7 High Light 101.8 20% Glycerol 102.4 b No Dye 101.7 Normal Light 101.6 Deionised Water 101.0 a

LSD (p=0.05)

NS

NS

0.45

# Values are percentages

64

Table 6.3: Means for the effect of the interaction between solution type and light intensity on solution uptake rate by foliage stems of E. robusta

Solution Light Intensity

High Light Normal Light 20% Glycerol #0.86 bc 0.73 ab

Deionised Water 1.14 c 0.46 a LSD: (p=0.05) 0.29

# Values are gg-1 fw d-1 Table 6.4: Means for the effect of the interaction between dye treatment and light intensity on relative mass of foliage stems of E. robusta

Light Intensity Dye Treatment Dye No Dye

High Light #102.1 b 101.5 ab Normal Light 101.3 a 101.9 ab

LSD: (p=0.05) 0.64 # Values are percentages Table 6.5: Effect of dye treatment, light intensity and solution type on flourescence scores for foliage stems of E. robusta

Dye Treatment

F Score Light Intensity F Score Solution F score

Dye #3.38 b High Light 2.29 20% Glycerol 2.79 b No Dye 1.00 a Normal Light 2.08 Deionised Water 1.58 a

LSD (p=0.05)

0.34

NS

0.34

# Scored on a 1-5 scale Movement of glycerol solution The solution type did not influence the uptake of solution over the 3 days that the experiment was conducted (Figures 6.1 and 6.2). However the solution type seemed to influence the degree of fluorescence observed on the leaf blade and in the mid-rib (Figure 6.3). Florescence scores taken on leaves treated with the 20% glycerol solution were much high than for those on leaves treated with deionised water. Within treatment solution type, there was little or no difference in the degree of fluorescence of leaf blades, irrespective of the leaf’s position on the stem or the time at which the dye was added to the uptake solution. In all cases, the mid rib gave a much higher fluorescence score than the leaf blade. As with intensity of fluorescence observed in the leaves, the day of adding the dye did not influence the fluorescence intensity found in the stem segments (Figure 6.4). However, unlike leaf position, there were differences in the intensity of fluorescence between the different stem segments. Whilst all segments had a relatively high fluorescence intensity, it was highest for those segments nearer the base of the stem.

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Figure 6.1 : Effect of day of application of apoplastic dye (0.1% pyranine solution) on solution uptake by stems of E. robusta in deionised water

Figure 6.2: Effect of day of application of apoplastic dye (0.1% pyranine solution) on solution uptake by stems of E. robusta in 20% glycerol solution.

0

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DYEPRESENT

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DYEPRESENT

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66

Figure 6.3: Effect of time of application of apoplastic dye (0.1% pyranine solution) on intensity of colour of leaves at various ascending canopy levels in E. robusta.

Figure 6.4: Effect of time of application of apoplastic dye (0.1% pyranine solution) on intensity of colour of foliage stems in ascending zones in E..robusta.

0

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Day 1 Day 2 Day 3 Day 1 Day 2 Day 3

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re Leaf 1Leaf 2Leaf 3Leaf 4Leaf 5Midrib

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20-25 cm

25-30 cm

- Glycerol + Glycerol

67

Discussion of results Effect of dye on the uptake of glycerol solution This experiment shows that the presence of the fluorescent dye, pyranine, does not have any effect on the uptake of glycerol solution by cut stems of E. robusta. Pyranine can therefore be used to assist in studying the movement of glycerol solution through such cut stems. It was interesting to note that stems that took up the same amount of deionised water as those that took up glycerol solution, both with added dye, received very different fluorescence scores. This suggests that the glycerol alters the perception of the degree of fluorescence. This may be due to the shine the glycerol creates on the surface of the leaf thereby increasing the degree of reflection. For this reason the degree of fluorescence caused by the presence of pyranine cannot be used as a measure of the amount of solution taken up, unless it is standardised for each solution type. Movement of glycerol solution This experiment supported the above finding that the uptake solution can alter the perceived fluorescence. It also suggests that, at least during a 3 day period of uptake, the glycerol solution is continually and evenly distributed to all leaves on the stem. The reduced intensity of fluorescence in the upper regions of the stem tissue is likely to be due to the limited apoplastic spaces within this immature tissue.

Implications Pyranine dye can be used to assist in studying the movement of glycerol solution through cut stems of E. robusta. The degree of fluorescence caused by pyranine dye cannot be used to measure of the amount of solution taken up by cut stems of E. robusta, unless it is standardised for each solution type. Based on the location of fluorescence in the stem and leaf tissues, glycerol moves relatively quickly and is distributed fairly evenly throughout these tissues over the initial 3 day period. In fact after as little as 24 hours of uptake of glycerol solution, the solution had been distributed to the peripheral leaves.

Recommendations To characterise the movement of glycerol or any other uptake solution in a stem of E. robusta, repeat measurements are required and should be based on a shorter observation period from the time when the dye is added to the uptake solution. Further experimentation is required to determine whether sufficient glycerol can accumulated in all areas of the stem inside the 3 day period and still maintain the qualities expected of an uptake preservation solution in terms of foliage suppleness and colour

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7. Do plant growth regulators affect the uptake of glycerol solution by Eucalyptus robusta? Introduction Plant growth regulators and other chemicals are known to influence stomatal behaviour. As a consequence, they influence the rate of transpiration and, thereby, the rate of solution uptake. In particular, the effect of the plant growth hormone, abscisic acid (ABA), on stomatal behaviour is well documented. ABA prevents stomatal opening and causes stomatal closure in virtually all plant species investigated (Willmer, 1983). When the level of ABA within leaves increases, a number of stomatal guard cell activities are almost simultaneously set in motion. These include a decrease in guard cell malate levels, an increase in the levels of starch in guard cells, and/or the movement of potassium ions (K+) out from within the guard cells. A number of phytotoxins produced by plant pathogens also have an effect on stomatal behaviour, either by promoting or inhibiting their opening. Fusicoccin, a phytotoxin produced by submerged cultures of the fungal pathogen Fusicoccum amygdali has been reported to cause opening of stomata when applied to leaves or epidermal strips. It can also overcome the ‘closing effects’ of ABA. Fusicoccin influences ion transport across the plasmalemma, increasing the extrusion of hydrogen ions (H+) from the guard cells and increasing the uptake of K+. This chapter investigates the effect of ABA and fusicoccin on the uptake of glycerol solution by cut stems of Eucalyptus robusta.

Objectives 1. Investigate the effect of ABA and fusicoccin applied as a pre-treatment pulse on the uptake of

glycerol solution by foliage stems of Eucalyptus robusta. 2. Investigate the effect of ABA and fusicoccin added to the uptake solutions on the uptake of

glycerol solution by foliage stems of Eucalyptus robusta.

Methodology Investigating the effect of pre-treatment pulse application of plant growth regulators Stems of E. robusta foliage were harvested from a managed garden at The University of Queensland Gatton and taken to the laboratory while standing into a solution and covered with plastic film. Harvested stems were re-cut to 10 cm in length, each with 2 leaves. They were pulsed for 1 day in either a solution of abscisic acid (50 µM ABA), fusicoccin (10 µM) or deionised water, as a control. Pulsing was conducted in a postharvest laboratory at 20°C and 70-80% RH and a light intensity of 60-70 µEinm-2s-1. After pulsing, stems were transferred to individual vases containing one or other of the two treatment solutions; 20% glycerol (v/v) as a preservative solution and deionised water as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of these solutions. Three replicates, each consisting of a cut stem in its own vase, were set up in a completely randomised design under the conditions which had prevailed during the pre-treatment. Solution uptake was recorded at 6, 12, 24, 30 and 48 h and daily thereafter up to 5 days. Investigating the effect of application of plant growth regulators to uptake solution Stems of E. robusta foliage were harvested from a managed garden at The University of Queensland Gatton. They were taken to the laboratory while standing in a solution and covered with plastic film. Harvested stems were re-cut to 20 cm, each with 2 leaves. These were placed into individual vases

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containing a combination of a plant growth regulator treatment and one of the two uptake solutions. The plant growth regulator treatments were abscisic acid (10 µM ABA), fusicoccin (10 µM) or deionised water, as a control. The uptake solution treatments were 20% glycerol (v/v) as a preservative solution and deionised water as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the uptake solutions. Five replicates were placed in a controlled environment at 24 ±1°C and 65 ±5% RH during the experiment. A completely randomised design was used. Solution uptake was recorded daily for 5 days. Observations were noted of leaf colour changes from green to brown, as an indicator of glycerol accumulation. Calculations for rate of uptake, the amount of solution taken up and the relative change in mass For each assessment, the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time in hours. A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results pertaining to this experiment can be found in Appendix 8.

Detailed results Effect of pre-treatment pulse application of plant growth regulators Pulsing cut stems of E. robusta with the plant growth regulators only had a significant effect on the uptake rate of the solutions within the first 12 hours of their being transferred to the uptake solutions (Table 7.1). At this time, uptake was fastest by those stems treated with fusicoccin, irrespective of the type of uptake solution (Figures 7.1(a) and 7.1(b)). There was no significant difference between the pre-treatment pulse in ABA and deionised water at 12 hours. However, uptake tended to be slowest in those stems pre-treated with ABA. This trend continued throughout the experimental period for the stems in the uptake solution of deionised water (Figure 7.1(a). There was no obvious trend for the uptake rate of stems in the glycerol solution (Figure 7.1(b). Overall, there was no difference in the rate of uptake of the two treatment solutions until Day 5. At Day 5 the uptake rate for the glycerol solution was only 0.003 gg-1fw h-1, while the uptake rate was about 10 times faster for deionised water at 0.029 gg-1fw h-1.

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The amount of uptake solution accumulated by the stems was only influenced by the pre-treatment pulse at the 6 h and 12 h observations. At the 12 h observation, stems treated with deionised water and fusicoccin had taken up an average of 0.241 gg-1fw h-1 while those which had been pre-treated with ABA had only accumulated about half that amount (0.127 gg-1fw h-1). The uptake solution type was found not to significantly affect cumulative solution uptake until later in the experimental period. Whilst there was no interaction between the uptake solution type and the hormone pre-treatment pulses, fusicoccin treated stems in the deinoised water uptake solution accumulated more solution than the control. These, in turn, accumulated more than the ABA treated stems (Figure 7.2(a)). Despite the large drop in relative mass of the ABA pre-treated stems transferred to deionised water uptake solution (Figure 7.3(a)), there were no significant effects of the hormone pre-treatments or the uptake solutions on the change in relative mass of the stems. Table7.1: Effect over time of hormone pre-treatment on rate of solution uptake by stems of E. robusta. Means followed by the same letter are not significantly different at p=0.05.

Treatment 6 h 12 h 24 h 48 h 72 h 96 h 120 h No Hormone 0.0165 b #0.0190 a 0.0181 0.0138 0.0151 0.0155 0.0125 Fusicoccin 0.0187 b 0.0258 b 0.0195 0.0166 0.0216 0.0215 0.0186

ABA 0.0093a 0.0119 a 0.0195 0.0121 0.0152 0.0120 0.0138 LSD

p=0.05 0.0062 0.0082 NS NS NS NS NS

# gg-1 fw h-1

71

Figure 7.1(a): Effect of hormone pre-treatment on rate of solution uptake by stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.1(b): Effect of hormone pre-treatment on rate of solution uptake by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol solution i.e. control; ν = fusicoccin; σ = ABA). No bars indicate non-significance.

0

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h

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e (g

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)

- Glycerol

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Figure 7.2(a): Effect of hormone pre-treatment on cumulative solution uptake by stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.2 (b): Effect of hormone pre-treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol solution i.e. control; ν = fusicoccin; σ = ABA). No bars indicate non-significance.

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Figure 7.3(a): Effect of hormone pre-treatment on relative mass of stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.3(b): Effect of hormone pre-treatment on relative mass of stems of E. robusta in 20% glycerol solution (υ = glycerol solution i.e. control; ν = fusicoccin; σ = ABA). No bars indicate non-significance.

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Effect of application of plant growth regulators to uptake solution Initially there was no significant difference in the solution uptake rate between the glycerol solution and the deionised water. Across both solutions, however, solution uptake rate was significantly less in the presence of ABA (average 0.935 gg-1fw d-1) than when fusicoccin or no hormone were added (1.377 or 1.167 gg-1fw d-1, respectively). Throughout the remainder of the experiment, it was observed that solution uptake was significantly faster in the presence of fusicoccin, followed by the no hormone treatment (except for Day 3) (Table 7.2). Uptake rate of glycerol solution dropped dramatically from an average across hormone treatments of 1.203 gg-1fw d-1 at Day 1 to 0.193 gg-1fw d-1 at Day 2. Glycerol solution uptake remained slow for the rest of the experimental period (Figure 7.4(b)). In contrast, uptake of deionised water increased to 1.636 gg-1fw d-1 on Day 2 from 1.117 gg-1fw d-1 on Day 1. The addition of fusicoccin significantly improved the uptake of deionised water compared with the addition of no hormone at all or ABA (Figure 7.4(a)). The increase in cumulative uptake of glycerol solution was minimal during the experiment, with the hormone treatments having no significant effect (Figure 7.5(b)). Approximately twice as much deionised water was accumulated when fusicoccin was added to the uptake solution as for the other treatments (Figure 7.5(a)). Relative mass ranged from 96.1 to 107.3% over the experimental period (Figures 7.6(a) and 7.6(b)). Although overall the addition of hormones to the uptake solutions did not have a significant effect on the relative mass of E. robusta stems, there were differences between the uptake solutions themselves. With the exception of Day 1, the relative mass of those stems taking up the glycerol solution was greater (average for days 2-5, 104.7%) than for stems in deionised water (average for days 2-5, 100.8%), due to the mass of glycerol compound in the stems. Table 7.2: Effect over time on rate of solution uptake by stems of E. robusta pulsed with hormones. Means followed by the same letter are not significantly different at p=0.05.

Day 1 Day 2 Day 3 Day 4 Day 5 No Hormone #1.1668 b 0.6925 a 0.7374 0.5884 a 0.6725 b Fusicoccin 1.3773 b 1.2992 b 1.0106 1.1032 b 1.1871 c

ABA 0.9351 a 0.7570 a 0.7059 0.4180 a 0.4180 a LSD (p = 0.05) 0.2204 0.2749 NS 0.3455 0.2539 # gg-1 fw d-1

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Figure 7.4(a): Effect over time of hormone treatment on solution uptake rate by stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.4(b): Effect over time of hormone treatment on solution uptake rate by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol solution i.e. control; ν = fusicoccin; σ = ABA). Bars show LSD (p=0.05) between means for the interaction of hormone treatment and solution and no bars indicate non-significance.

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Figure 7.5(a): Effect over time of hormone treatment on cumulative solution uptake by stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.5(b): Effect over time of hormone treatment on cumulative solution uptake by stems of E. robusta in 20% glycerol solution (υ = 20% glycerol solution i.e. control; ν = fusicoccin; σ = ABA). Bars repshow LSD (p=0.05) between means for the interaction of hormone treatment and solution and no bars indicate non-significance.

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Figure 7.6(a): Effect over time of hormone treatment on relative mass of stems of E. robusta in deionised water (υ = deionised water i.e. control; ν = fusicoccin; σ = ABA).

Figure 7.6(b): Effect over time of hormone treatment on relative mass of stems of E. robusta in 20% glycerol solution (υ = 20% glycerol solution i.e. control; ν = fusicoccin; σ = ABA). Bars show LSD (p=0.05) between means for the interaction of hormone treatment and solution and no bars indicate non-significance.

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Discussion of results Effect of pre-treatment pulse application of plant growth regulators The technique of pre-treating cut stems of E. robusta with a 1-day pulse of either ABA or fusucoccin did not affect the uptake of glycerol solution. It was expected that the pre-treatment pulse with ABA would decrease solution uptake by causing closure of the stomata and, thereby, reducing transpiration. In contrast, it was expected that pre-treatment with fusicoccin would result in an increase in the uptake of the glycerol as the stomata opened and transpiration increased. Neither of these two pre-treatments altered solution uptake compared with the control stems pulsed with deionised water. Effect of application of plant growth regulators to uptake solutions The uptake of glycerol was not enhanced by the addition of fussicoccin when compared to the control of not adding hormone. The addition of ABA, in contrast, tended to reduce the uptake of the glycerol solution.

Implications The plant growth regulators, ABA and fusicoccin, at the concentrations tested, should not be used to pre-treat cut stems of E. robusta for uptake preservation with glycerol solution. These plant growth regulators do not provide any benefits.

Recommendations Further experimentation is required to determine whether the use of these plant growth regulators may provide any benefit to the uptake preservation process for E. robusta. Such experimentation should consider varying the duration of the pulse, varying the concentration of the plant growth regulators in the pulsing or uptake solution, the direct application as a spray or dip to plant foliage, as well as the use of alternative chemicals that control stomatal aperture. Moreover, testing may need to be done on a species by species basis.

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8. Does the uptake of glycerol solution affect the respiration of cut Eucalyptus robusta and E. tetragona? Introduction Respiration is a reflection of metabolic activity. That is, the greater the overall metabolic activity of a given tissue, the higher is its respiration rate (Taiz and Zeiger, 1991). Glycerol is used by the cut flower and foliage industry to preserve plant material. Its physiological effects have not, however, been described. Exposure of plant tissue to polyols, such as mannitol, glycerol and polyethylene glycol have been reported to slow down respiration and synthesis processes (Greenway and Leahy, 1970). Preliminary studies with Eucalyptus cinerea cut foliage treated with 30% glycerol revealed a decline in respiration rate within half a day of commencing the treatment (Campbell, 1996). This chapter aims to describe the physiological effects of glycerol on E. robusta and E. tetgragona.

Objectives 1. Determine the effect of glycerining on the respiration rate of cut foliage stems of Eucalyptus

robusta and E. tetragona. 2. Compare the glycerol solution uptake by Eucalyptus robusta and E. tetragona subjected to the

same treatment.

Methodology Stems of Eucalyptus robusta foliage were harvested from a managed garden at The University of Queensland, Gatton on the morning that the experiment was to commence. On return to the laboratory, stem ends were re-cut under water to 25 cm in length, with 5 leaves. Foliage stems of E. tetragona were harvested from a commercial plantation known as “Brookvale Park” near Cecil Plains in the afternoon of the day prior to commencing the experiment. On return to the laboratory, each stem was re-cut under water to 20 cm in length, with 15 leaves. For each of the experiments, two treatment solutions were used: 20% glycerol (v/v) as a preservative solution and deionised water as a control. A biocide of 10 mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Ten replicates, each consisting of a cut stem in its own vase, were placed in an access area at 20°C and 62-70% RH under continuous light of 2.5-3 µEinm-2s-1 and air speed of less than 1 ms-1 during experimentation. A completely randomised design was used.

Respiration was measured by sealing each cut stem in its treatment vase in a tall 3.2 L capacity plastic respiration container. After about 1 hour, a 1 mL sample of headspace gas was collected by inserting the needle of a hypodermic syringe into the rubber sampling port of the respiration containers. The gas sample was immediately injected into a gas chromatograph equipped with a TCD detector. The amount of CO2 produced by each stem was expressed as mL CO2 L-1h-1.

Respiration rate and solution uptake by stems of E. robusta were measured at 6, 12, 24, 36 and 48 hours, daily thereafter to Day 7 and then at Day 9 for the respiration rate and at Day 9.5 for the solution uptake recording. Similar measurements were taken for E. tetragona at 12, 24, 42 and 48 hours and daily thereafter to Day 7. For each assessment, the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae:

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Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours, and n-1 is the previous assessment time in hours. A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results pertaining to this chapter can be found in the Appendix 9.

Detailed results Solution uptake E. robusta The solution uptake rate by the foliage stems of E. robusta in the glycerol solution at 6 hours was higher (0.084 gg-1 fw h-1) than for those stems in the deionised water (0.066 gg-1 fw h-1) (Figure 8.1). Uptake of glycerol solution then dropped dramatically to 0.003 gg-1 fw h-1 at Day 2 and remained at a very low level for the remainder of the experiment. In contrast, deionised water uptake fluctuated for the first 3 days of the experiment and then steadied at around 0.004 gg-1 fw h-1 before declining. The cumulative uptake of the glycerol solution showed a constant uptake of solution for the first 36 hours to 1.24 gg-1 fw (Figure 8.2). However, after this time, uptake slowed considerably. Over the next 7.5 days, these stems only accumulated an additional 0.19 gg-1 fw. By the same time, those stems in the deionised water had accumulated a total of 8.87 gg-1 fw of solution by steady uptake throughout the experiment. The transpiration rate at 6 hours was 0.089 and 0.065 gg-1 fw h-1 for glycerol solution and deionised water respectively, and thus similar to their respective solution uptake rates. The similarity between transpiration rate and solution uptake rate continued throughout the experimental period (Figure 8.3). The relative mass of the deionised water treated stems was also relatively steady during the experimental period, averaging around 100.7% and deviating by no more than 0.3% (Figure 8.4). The situation was quite different for the glycerol treated stems. The relative mass of these stems rose from 96.8% to 104.4% in the first 24 hours and was stable at this level for the next 6 days. However, by Day 9.5, relative mass had risen again reaching 113.0%. E. tetragona E. tetragona had a very slow solution uptake rate compared to E. robusta, irrespective of uptake solution type (Figure 8.5). That is, even the uptake of deionised water was very slow, fluctuating within a range of 0.004 and 0.010 gg-1 fw h-1. In contrast, uptake of the 20% glycerol solution peaked at 0.021 gg-1 fw h-1 at 48 hours and then gradually declined to 0.001 gg-1 fw h-1 by the completion of the experiment.

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Cumulative uptake of solution also produced unexpected results. There was no difference in the amount of solution taken up by the stems of E. tetragona in either deionised water or the glycerol solution during the first 24 hours, nor was there a difference for the last 2 recording times on Days 6 and 7 (Figure 8.6). During the bulk of the experiment period solution uptake was greatest by those stems in the glycerol solution. By Day 3 these stems had accumulated almost twice as much solution as those in deionised water. At the completion of the experiment, the data suggest that uptake of glycerol solution was starting to plateau, while uptake of deionised water was continuing to increase. The transpiration rates for both the deionised water and glycerol solution corresponded with their respective uptake rates and thus were very low, the maximum rate of 0.022 gg-1 fw h-1 for the glycerol solution being recorded at 48 hours and that for deionised water (0.0095gg-1 fw h-1) recorded at 144 hours (Figure 8.7). During the first 42 hours of the experiment, the relative mass of stems increased and there was little difference between the treatment solutions (Figure 8.8). After this time, stems in deionised water remained relatively steady, rising from 101.3% to 101.9% by Day 7. In contrast, the relative mass of stems treated with glycerol dropped to 96.8% at Day 5 before recovering to the same level as those stems in deionised water. From 42 hours, leaves on stems in the glycerol solution started to show discolouration, while those in deionised water remained green. By completion of the experiment, all stems in glycerol solutions had in excess of 75% leaf browning. This discolouration tended to be concentrated around the mid-rib and along the veins, with the leaf margins often remaining green. Respiration rate E. robusta Cut stems of E. robusta held in DI water had relatively constant respiration rates ranging from 113-180 mL CO2 kg -1 fw h-1 during the 7-day postharvest period (Figure 8.9). Stems in 20% glycerol solution had similar initial respiration rate to those in DI water. Their respiration rate then steadily declined to about 25 mL CO2 kg-1 fw h-1 by Day 6. E. tetragona E. tetragona stems in deionised water and glycerol solution initially had high respiration rates of 163 and 212 ml CO2 kg-1 fw h-1, respectively (Figure 8.10). Throughout the experiment, stems in both solutions showed a similar trend of declining respiration rates over time. The trend, however, was most rapid during the first 3 days for those stems in the glycerol solution. At Day 3, the respiration rate of these stems dropped below the level of the stems in deionised water, to 93 ml CO2 kg-1 fw h-1. Stems in deionised water had a steady decline in respiration rate during the experiment period, to a low of 83 ml CO2 kg-1 fw h-1 on Day 7. It is interesting to note differences in respiration rates of E. tetragona stems of differing maturities. Some of the stems used in this experiment had mature foliage and fruits were often present, whilst others had no fruits and the foliage was immature. At 42 hours there was a tendency for those stems with immature foliage to have a higher respiration rate than those with mature foliage (Table 8.1). This was most marked in those stems treated with the glycerol solution. Table 8.1 Respiration rate of E. tetragona stems of differing maturities after 42 hours of uptake of preservative solutions. Respiration rate (ml CO2 kg-1 fw h-1)

Immature stems (number) Mature stems (number) 20% Glycerol solution 206.7 (6) 112.4 (4) Deionised water (control) 180.9 (4) 118.2 (6)

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Figure 8.1 Effect over time of solution type on rate of solution uptake by foliage stems of E. robusta (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

Figure 8.2: Effect of solution type on cumulative solution uptake by foliage stems of E. robusta (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

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Figure 8.3: Effect over time of solution type on transpiration rate of foliage stems of E. robusta(υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

Figure 8.4: Effect over time of solution type on relative mass of foliage stems of E. robusta (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

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Figure 8.5: Effect over time of solution type on rate of solution uptake by foliage stems of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

Figure 8.6: Effect of solution type on cumulative solution uptake by foliage stems of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

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Figure 8.7: Effect over time of solution type on transpiration rate of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

Figure 8.8: Effect over time of solution type on relative mass of foliage stems of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

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Figure 8.9: Effect over time of solution type on transpiration rate of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

Figure 8.10: Effect over time of solution type on respiration rate of foliage stems of E. tetragona (υ = deionised water; ν = 20% glycerol solution). Bars show LSD (p=0.05) between means.

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Discussion of results Solution uptake characteristics of foliage stems varied markedly between E. robusta and E. tetragona, despite some similarites in respiration, especially in deionised water treatments. However, E. tetragona had considerably higher respiration rates than E. robusta in glycerol solution until 48 hours. Thereafter its rates declined and were similar to those of E. robusta. Despite this similarity, solution uptake was much slower in E. tetragona than in E. robusta, with the average across solution treatments at 12 hours showing an 8-fold difference. Ultimately the two species took up similar amounts of glycerol solution. This, however, was achieved by E. robusta after only 36 hours, whilst some 168 hours were required for E. tetragona. The results show that this difference in uptake rate was related to differences in transpiration rates, which could be attributed to differences in cuticle thickness between these species (Chapter 2). The cuticle thickeness of E. tetrgona was about 3.5 times the thickness of E. robusta on both leaf surfaces. There were some similarilites between these results and those obtained for the experiment in Chapter 2. Culmulative uptake of glycerol by E. tetragona was lower than that for E. robusta for the first 2 days and thereafter they were similar. It is possible that differences noted in uptake are related to differences in morphology and anatomy of the two species (Chapter 2). However the physiological status of the plant (due to climatic differences) and its postharvest handling procedures may have an even great influence.

Implications Species of Eucalyptus differ in their ability to take up glycerol. Thus, there may be a need to change the length of the preservative process on a per species basis. Under the same environmental conditions, E. tetragona requires more time than E. robusta to take up the desired amount of solution and, therefore, of glycerol.

Recommendations Modify preservative protocols for different species.

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9. Is the uptake of preservative solution by stenanthemum affected by the type of humectant used and its concentration? Introduction Stenanthemum scortechenii (syn. Cryptandra scortechenii) is known in the cut flower trade as stenanthemum and ‘Corroboree flower’. It is a woody perennial species that produces its flowers in ‘pom-pom’-like inflorescences along the length of its stems during winter and spring. David Hockings of Maleny selected this Queensland native species for its vase life, ease of cultivation and suitable habit in terms of display of flowers. In addition, stenanthemum can be sold either as a fresh or dried cut flower, in its natural state or dyed. As with many species, the stenanthemum stems and foliage become quite brittle on air drying. Thus, there is potential to improve its postharvest quality for the dried flower market by developing appropriate treatments for the preservation of this new cut flower crop. This chapter investigates the uptake of humectants by stenanthemum.

Objectives 1. Investigate the effect of glycerol concentration on uptake of solution by stenanthemum. 2. Investigate the effect of humectant type and concentration on uptake of solution by

stenanthemum.

Methodology General methodology for investigating the rate of uptake, the amount of solution taken up and the relative change in mass Flowering stems of stenanthemum cv. B18 were harvested from Maleny, Queensland. The stems were transported within 3 hours in a polystyrene esky to The University of Queensland, Gatton for experimentation. Upon arrival, stems were placed in water for 2 to 3 hours to re-hydrate. Immediately before experimentation commenced, stem ends were re-cut under water to 30cm in length and weighed. Ten replicates per treatment, each consisting of a cut stem in its own vase, were placed in a controlled environment at 24 ±1°C and 65 ±5% RH. Experimentation involved using a completely randomised design. Solution uptake and stem weight were recorded once daily over 7 days. Two separate experiments were conducted, one relating to the concentration of glycerol solution and the other investigating different humectants, as outlined below. For each assessment the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0

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Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where, t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time in hours. To calculate the actual amount of glycerol taken up, the total weight of solution taken up was multiplied by a factor based on the glycerol concentration and the specific gravity of glycerol (1.262). Thus for a 10% glycerol solution (v/v), the factor is 0.123, that is it is equivalent to a 12.3% glycerol solution (w/w). For a 20% glycerol solution (v/v), the factor is 0.240, and 0.351 for a 30% solution (v/v). A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results can be found in Appendix 10. Modifications for investigating the effect of glycerol concentration Three glycerol treatment solutions were used:10%, 20% and 30% glycerol (with concentrations determined on a volume to volume basis), with deionised water used as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Modifications for investigating the effect of humectant type and concentration Four humectant treatment solutions were used: 20% glycerol, 15% glycerol plus 5% polyethylene glycol (PEG), 10% glycerol plus 10% PEG and 20% PEG. All concentrations were on a volume to volume basis. Deionised water was used as a control treatment. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions.

Detailed results Effect of glycerol concentration This experiment showed that the uptake rate of deionised water by stenanthemum was significantly greater than the uptake rate of the glycerol solutions (Figure 9.1). Whilst there was an overall trend that the uptake rate decreased with increasing glycerol concentration, there was rarely any significant difference between the three concentrations. Across the 3 glycerol treatments, the mean initial uptake rate was 0.018 gg-1fw h-1. However, this declined during the experimental period to 0.008 gg-1fw h-1 by Day 4 and 0.007 gg-1fw h-1 by Day 6. In contrast, over the same period, the uptake rate of deionised water was relatively constant at around 0.027 gg-1fw h-1. Cumulative uptake of the glycerol solutions by stenanthemum showed a similar pattern of decreasing accumulation with increasing concentration of glycerol (Figure 9.2). There was no significant difference between stems accumulating the 10% and 20% solutions. At Day 4 cumulative uptake was 1.582 gg-1fw and 1.324 gg-1fw, respectively, the average being 1.453 gg-1fw. However, this was significantly more than the cumulative uptake of the 30% solution, which at Day 4 was 0.752 gg-1fw. However, when the actual cumulative uptake of glycerol per se was calculated, this trend did not appear (Table 9.1). On Day 1, those stems in the 30% glycerol solution had taken up nearly 2.5 times the amount of glycerol as the stems in the 10% glycerol treatment, despite the fact that they had taken up less solution. From Day 2 the stems in the 20% solution recorded the greatest accumulation of glycerol, and from Day 4 this was significantly greater than that taken up by stems in either the 10% or 30% glycerol solutions.

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Table 9.1. Cumulative uptake of glycerol compared with total solution uptake by stenanthemum (gg-1

fw). Different letters within each column show significant differences in uptake for either total solutions or glycerol solutions (LSD p=0.05). Glycerol solution Uptake of: Day 1 Day 2 Day 4 Day 6

10% Solution 0.466 A 0.992 B 1.582 B 2.017 B

Glycerol 0.057 a 0.122 a 0.195 a 0.248 a

20% Solution 0.462 A 0.902 B 1.324 B 1.568 B

Glycerol 0.111 b 0.216 b 0.318 c 0.376 c

30% Solution 0.384 A 0.563 A 0.752 A 0.872 A

Glycerol 0.135 b 0.198 b 0.264 b 0.306 b There was little difference between the glycerol treatments throughout the experiment with respect to the relative mass of the stenanthemum stems (Figure 9.3). The average relative mass was 95% for these stems, with a range from 90.0% to 99.8%. The stems treated with deionised water had an initial relative mass of 97.9%, which was not significantly different from that of the stems in any of the glycerol solutions. However, as the experiment progressed, they showed a steady decline in relative mass.

Figure 9.1: Effect over time of glycerol concentration on solution uptake rate by foliage stems of Stenanthemum sp. (υ = deionised water control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) between means for significant effects.

0

1

2

3

4

5

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Cum

ulat

ive

Solu

tion

Upt

ake

(gg-1

fw)

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Figure 9.2: Effect of glycerol concentration on cumulative solution uptake by foliage stems of Stenanthemum sp. (υ = deionised water control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) between means for significant effects.

Figure 9.3: Effect over time of glycerol concentration on relative mass of foliage stems of Stenanthemum sp. (υ = deionised water control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol). Bars show LSD (p=0.05) between means for significant effects.

0

0.01

0.02

0.03

0.04

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Solu

tion

Upt

ake

Rat

e (g

g-1

fw h

-1)

60

70

80

90

100

110

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Rel

ativ

e M

ass

(%)

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Effect of humectant type and concentration The uptake rate of the humectant solutions was significantly slower than that of the control treatment of deionised water (Figure 9.4). In this experiment, humectant type and concentration did not influence the uptake of solution by stenanthemum. The average uptake rate of the humectant solutions at Day 1 was 0.010 gg-1 fw h-1. However, by Day 4 this had dropped to 0.003 gg-1 fw h-1, and remained around this level for the rest of the trial period. There was a general increase in the cumulative uptake of solutions throughout the experimental period (Figure 9.5). The greatest amount of solution taken up was by those stems in the control treatment, although at Day 1 there was no difference between any of the treatments. Cumulative uptake of all of the humectant treatments was very low at less than 0.9 gg-1 fw and there was no difference between the treatments. The relative mass of all stems was quite low on Day 1 (73 to 84%) and continued to decline throughout the experimental period (Figure 9.6). There was no significant difference between any of the treatments.

Figure 9.4: Effect over time of humectant type on rate of solution uptake by stems of Stenanthemum sp. (υ = deionised water control; ν = 20% PEG; σ = 5% PEG + 15% Glycerol; λ = 20% Glycerol; = 10% PEG +10% Glycerol. Bars show LSD (p=0.05) between means for significant effects

0

0.005

0.01

0.015

0.02

d 1 d 2 d 3 d 4 d 5 d 6 d 7

Time

Solu

tion

Upt

ake

Rat

e (g

g-1 fw

h-1

)

93

.

Figure 9.5: Effect of humectant type on cumulative solution uptake by stems of Stenanthemum sp. (υ = deionised water control; ν = 20% PEG; σ = 5% PEG + 15% Glycerol; λ = 20% Glycerol; = 10% PEG +10% Glycerol. Bars show LSD (p=0.05) between means for significant effects.

Figure 9.6: Effect over time of humectant type on relative mass of stems of Stenanthemum sp. (υ = deionised water control; ν = 20% PEG; σ = 5% PEG + 15% Glycerol; λ = 20% Glycerol; = 10% PEG +10% Glycerol). No bars indicate non-significance.

0

0.5

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(gg-1

fw)

60

70

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90

100

d 1 d 2 d 3 d 4 d 5 d 6 d 7

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ass

(%)

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Plate 9.1 Transverse section of flowering stem of Stenanthemum scortechenii showing the many small xylem vessels.

Plate 9.2 Transverse section of Eucalyptus robusta showing xylem vessels which were typically larger than those for Stenanthemum scortechenii.

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Discussion of results Effect of glycerol concentration This experiment illustrates that whilst weaker solutions of glycerol may be taken up faster by stems of stenanthemum, actual accumulation of glycerol can be more rapid at higher concentrations. Overall, the uptake rate was very low compared to some of the eucalypt species used in this study. It is possible that this relates to the very small xylem vessels in stenanthemum (Plates 9.1 and 9.2). The leaves of stenanthemum are adapted to minimise water loss by having small hairs on the underside of the leaves (Chapter 2, Plate 2.1). This however, may also reduce the uptake of solution by decreasing the rate of transpiration. In considering the relative mass of the stems during the experiment, the steady decline demonstrated by the stems in deionised water suggests that they were unable to continue to take up the water at the same rate as they were losing it through evapo-transpiration. This resulted in a loss of quality by these stems. In contrast, the humectant property of the glycerol, in this case irrespective of the solution concentration, was able to maintain the relative mass. Effect of humectant type and concentration Solution uptake by stems of stenanthemum was less in this experiment than in that investigating glycerol concentration, when common treatment solutions are compared (i.e. 20% glycerol solution and deionised water control). The reason for this is unclear. However it may reflect differences in the water status of plants in the field, prevailing environmental conditions and/or conditions during handling and transportation. The experiment also did not test whether these low levels of solution uptake were sufficient to provide adequate preservation.

Implications For stenanthemum, it is possible to increase the concentration of the glycerol solution used and thereby reduce the time required to take up the required amount of glycerol. It is unclear whether polyethylene glycol alone or in combination with glycerol can be taken up by stenanthemum to a suitable level to provide any preservation of quality of the plant material.

Recommendations A 20% glycerol solution gives the most rapid accumulation of glycerol in stems of stenanthemum. Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of stenanthemum. Trials also are required to identify if there is an upper limit to the concentration of the glycerol solution that can be used to satisfactorily preserve stenanthemum. Further experimentation is required on fresh material of stenanthemum to determine whether polyethylene glycol alone or in combination with glycerol is a suitable humectant for preservation of this cut flower.

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10. Does relative humidity and type of packaging during storage effect the outturn quality of glycerined stenanthemum cut flowers? Introduction While floricultural exports of preserved material are increasing, the quality of preserved products is highly variable (Joyce 1997). Many factors influence the efficiency of uptake preservation (Dubois and Joyce 1992). However little is known of how these factors may influence the maintenance of product quality during packaging and marketing. In addition many different species are used as preserved products and are likely to perform differently. Plant materials preserved with humectants have a soft and supple texture, because they hold moisture. However, preserved material is susceptible to softening, sweating and microbial spoilage The amount of moisture held by a humectant in equilibrium with its environment depends on relative humidity (RH), temperature and the adsorption and desorption process (Bhandari, 1997). The adsorption and desorption properties of preserved products are influence by the RH of the surrounding air. The water holding capacity of most humectants increases steeply above 80% RH (Labuzza 1984). Fluctuations in temperature can cause condensation or dehydration (Bhandari, 1997) as well as encouraging the growth of micro-organisms. In addition, internal and external properties of the stem and leaves will influence the adsorption and desorption properties of the humectant. Packaging influences the microenvironment around preserved stems. Methods of controlling RH within the packages have been demonstrated by Shirazi and Cameron (1992). They sealed within low density polyethylene (LDPE) liners chemicals known to influence RH. Thus as LDPE has a low vapour transmission rate, RH within the package can be controlled independently of external RH (Shirazi and Cameron, 1992). This chapter looks at the effects of different humidity and packaging treatments during storage on the outturn quality of preserved stems of Stenanthemum scortechinii.

Objectives 1. Determine the effects of different relative humidities during storage on the outturn quality of

preserved Stenanthemum. 2. Evaluate different package regimes on the outturn quality of preserved Stenanthemum. 3. Develop a test to predict the stability of the preserved product under the variable conditions of

the consumers’ environment.

Methodology Stems of cut Stenanthemum preserved with 20% v/v glyercol solution were obtained from a commercial processor (David and Olive Hockings, Maleny Queensland). Stems were sorted for uniformity of visual appearance (colour) and touch (flexibility), and trimmed to a uniform length (30 cm). Lateral branches were removed from the lower 5 cm of the stem. Experiment 1 - Storage at different humidities Four stems were placed into each fibreboard box (33cm x 12cm x 5.5 cm) and five replicate boxes were placed into each of three storage treatments of 30, 60 and 90% relative humidity (RH). These storage treatments were achieved inside three 20 L plastic jars by passing an air flow rate of 1.36 mL sec-1 through calcium chloride solution (30% RH), water (90% RH) or an equal mixture of these flows (60% RH) (Figure 10.1 and Plates 10.1a and b).

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Figure 10.1 Relative humidity control system used for storage of stems of glycerined Stenanthemum. The stems in four boxes were used as replicates for determining the initial weight and monthly weight over a 6 month storage period from 9 October 1997 to 9 April 1998. Stems in the fifth box were the source of samples for an equilibrium relative humidity (ERH) test. The change in mass (% initial weight) was calculated each month. Observations were made on colour change, flexibility, sweating or mould development and leaf or flower drop.

Plate 10.1 Boxes of glycerined Stenanthemum stems inside 30% humidity jar (a) and those stems placed in the fibreboard carton (b).

Sat’d CaCl2

air

Low r.h.≅ 30%

Med. r.h.≅ 60%

High r.h.≅ 90%

dry air

air air

H2O

mixing

trap

trap

trapbubblingstone

needle valve

needle valve

oo:.o.

oooo..ooo.

……...

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Experiment 2 - Packaging conditions Five packaging treatments inside fibreboard boxes were evaluated; 1. Control - stems were placed in a fibreboard box with no chemical sachet or liner, 2. Stems were sealed in a low density polyethylene (LDPE) liner (35 µm thick), 3. Stems were sealed in a LDPE liner with a sachet of calcium chloride (CaCl2), to create a low

RH, ERH at 20°C being 32.3%. 4. Stems were sealed in a LDPE liner with a sachet of glucose (to create a moderate RH, ERH

at 20°C being about 60% (Bhandari 1997). 5. Stems were sealed in a LDPE liner with a sachet of potassium chloride (KCl), to create a

high RH, ERH at 20°C being 85.0%. There were 3 stems per bunch, 3 replicate bunches per liner and 3 liners per packaging treatment, one for each time of removal after 2, 4 and 6 months storage. The preserved flowers and chemical inside the LDPE liners were sealed by a heat sealing machine for ca. 7 seconds expelling as much air as possible. Stems of Stenanthemum in these packaging treatments were evaluated in two different storage environments, a packing shed, without temperature control and an air-conditioned room at 20°C and 95% RH. The change in mass (% initial weight) was calculated and ERH determined every 2 months for 6 months. Observations were made on colour change, flexibility, sweating or mould development and leaf or flower drop. Equilibrium relative humidity (ERH) test Joyce (1997) applied the ERH test to preserved plant material to evaluate its performance under different relative humidities. It is based on the fact that the RH of the headspace of an enclosed chamber containing a saturated salt is a direct and invariable function of the type of salt and the temperature. Equilibrium relative humidities have been accurately measured and tabulated against temperature (Solomon 1951, Rockland 1960, Weast 1977, Winston and Bates 1960). Useful salts for obtaining a range of relative humidities at 20°C are listed (Table 10.1). Table 10.1. Some salts that can be used to achieve a range of relative humidities (%) at 20°C (Shirazi and Cameron 1992, Rockland 1960). Salt Equilibrium relative

humidity (%) Saturation point

(g/20mL) Calcium chloride 32.3 17.4 Potassium carbonate dihydrate 44.0 23.5 Sodium hydrogen sulfate 52.0 6.1 Sodium nitrite 66.0 19.3 Sodium chloride 76.0 7.6 Ammonium chloride 79.5 8.2 Potassium chloride 85.0 7.4 Magnesium sulphate 90.0 7.8 Di-sodium hydrogen phosphate, 7 hydrate 95.0 3.4 The seven saturated salt solutions in Table 10.1 were prepared in 100 mL jars by dissolving, heating to 30°C, stirring and standing overnight at 20°C for precipitating of the excess salt. The plant tissue was supported above the salt solution on a plastic mesh stand (Plate 2). Initial ERH of Stenanthemum stems was determined by plotting the relative successive change in mass against relative humidities each week for 4 successive weeks. This exercise showed that a 1 week equilibrium period could be used. Relative change in mass (RM) was calculated as a % initial mass as follows;

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Difference between final weight (g) and initial weight (g) divided by initial weight (g) and multiplied by 100. ERH was recorded as the point on the regression between change in RM% and RH where the RM did not change (i.e. was 0).

Plate 10.2. Equilibrium relative humidity control system for preserved Stenanthemum stems. Statistical analysis A completely randomised design was used for both experiments. A split plot over time ANOVA was adopted to differentiate treatment means of the stems stored at 3 humidities using the balanced model procedure for Minitab Version 11 (Minitab, Inc., 1996). Relative humidity was used as the main unit factor with time of storage nested within each humidity. For the effects of different packaging, a standard balanced model, split plot over time ANOVA was used to analyse differences among treatment means. For the ERH test analysis, a hyperbolic model was fitted using the NLIN procedure in SAS for Windows TM Release 6.11 (SAS Institute Inc. 1996).

Detailed results Experiment 1 - Storage at different humidities The initial ERH of the Stenthanemum stems was 56%. Storage at 30% RH significantly reduced (p=0.05) the relative mass of stems by about 12 % during the first month of storage. Thereafter over the following five months, only a minor loss of relative mass occurred. However, after five months of storage at 30% RH this reduction in relative mass was significant (Table 10.2). Stems stored at 30% RH were brittle and some leaf and flower drop was observed after both the first and fifth month of storage. The ERH of tissue stored at 30% RH was reduced from an initial reading of 56% to about 40.0% after storage.

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Table 10.2: Relative mass (% initial weight) of glycerined Stenanthemum stems after storage at different humidities, together with least significant differences (n=4).

Storage Storage time (months)

humidity 1 2 3 4 5 Means

30% 88.1 88.1 87.9 87.8 86.7 88.7b

60% 100.4 97.2 99 5 99.4 99.2 99.1a

90% not included

in mean

126.0 - - - -

Means 94.3ab 93.7ab 93.6ab 93.0bc 92.6c

Main factors LSD 0.05 Humidity means 0.74 LSD 0.05 Storage time means 1.12 Interaction means LSD 0.05 Within same humidities 1.05 LSD 0.05 Between storage humidity by time 1.45

Stems stored at 60% RH showed a small but significant reduction in relative mass over the storage period, but were observed to retain their marketable quality. The ERH of stems stored at 60% RH increased slightly from 56% at the start of the storage period to 59% at the end of the experiment. Stems stored at 90% RH showed a large weight gain after one month of storage. These stems became sweaty, very soft and supported growth of fungal and bacterial organisms. These organisms were Cladosporium sp., Curvularia sp., Eurotium sp. Aspergillus sp. Chaetophoma sp. and Aureobasidium sp. Theses stems were discarded from the experiment after only one month of storage and thus were not included in the statistical analysis. Experiment 2 - Packaging conditions The packaging treatments used did not protect stems from deteriorating when they were stored at 20°C and > 90% RH. Stems in fibreboard cartons were mouldy within one month and all treatments were discarded by the fourth month of storage. Consequently results for this storage environment were not used in the analysis. Boxes stored in the packing shed (22-35°C) were less affected. Packaging treatments and storage times significantly (p=0.05) influenced the relative mass of Stenanthemum stems, and there was a significant interaction between packaging treatment and storage time (Table 10.3). After two months storage, stems held in the fibreboard carton had lost about 12 % of their inital weight, more than stems held in other packaging treatments.

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Table 10.3: Relative mass (% initial weight) of preserved Stenanthemum stems stored under different packaging conditions for 6 months in a packing shed (22-35°C), together with least significant differences (n=3). Means followed by the same letter within a storage period are not significantly different. Packaging treatment Removal from storage Packaging 2 months 4 months 6 months

treatment

means Fibreboard carton

88.2e 95.2b 95.1b 92.8d

Fibreboard carton and LDPE sealed liner 94.6c 93.9c 93.8b 94.1c

Fibreboard carton, LDPE sealed liner and CaCl2 sachet

90.3d 89.5d 89.4c 89.7e

Fibreboard carton, LDPE sealed liner and glucose sachet

95.3b 96.8a 96.8a 96.3b

Fibreboard carton, LDPE sealed liner and KCl

97.0a 97.8a 97.9a 97.6a

Storage time means 93.1b 94.6a 94.6a Main factors LSD 0.05 Packaging treatment means 1.12 LSD 0.05 Storage time means 0.48 Interaction means LSD 0.05 Within same main packaging treatments 1.06 LSD0.05 Between different packaging or storage time treatments 1.42

Stems in fibreboard boxes, with a sealed LDPE liner and a sachet of CaCl2 lost about 10% relative mass after two months storage. The ERH of these stems declined from an initial ERH of 56% to about 39% after storage. Leakage from the salt sachet spoilt the stems in this packaging treatment. Stems stored in fibreboard cartons with a sealed LDPE liner and a sachet of KCl showed the least reduction in relative mass after storage for 2 months and remained stable throughout the 6 month storage period. Similarly, stems stored in fibreboard cartons with a sealed LDPE liner and a sachet of glucose were not different to those stored with a KCl sachet after 4 and 6 months storage. These stems showed similar ERH at the end of the experiment viz.: KCl 52% and glucose 51%.

Discussion of results These experiments showed that the humidity of the storage environment affects outturn quality of preserved Stenanthemum stems. Storage at high RH (>80%) should be avoided due to the strong adsorption properties of humectants (Labuza 1984 and Bhandari 1997). Stems stored at 30% RH showed reduced relative mass during storage, and were susceptible to loss of quality by becoming brittle. Packing preserved stems into fibreboard containers without a LDPE liner means that they are susceptible to moisture loss or gain caused by RH of the external environment. Packaging treatments can be used to modify the RH within the fibreboard carton, and provide a stable environment for prolonged storage without quality loss. The sachets with glucose or KCl were both effective in maintaining ERH and outturn quality of product. The use of the desiccant CaCl2 salt reduced the outturn quality by excessive moisture loss from the product and spoilage. Quality management programs could be enhanced by adoption of the ERH test (Joyce 1997). It could be used to determine the suitability of a product to the environmental conditions in the consumers display area. Alternatively it could be used as a feedback loop to refine the preservation process.

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Implications Storage of preserved flowering stems at high relative humidity (>80%) and low relative humidity (30%) should be avoided as quality will be adversely affected by excessive adsorption of humectants or by moisture loss respectively. If stems are to be packed for prolonged storage, cartons should have a LDPE liner and a sachet of glucose or potassium chloride to maintain a stable environment and hence avoid loss of outturn quality. Adoption of the ERH test could determine the suitability of a product to the environmental conditions in the consumers display area. Alternatively it could be used as a feedback loop to refine the preservation process. Recommendations Growers need to consider the environment during the preservation process and storage to ensure outturn quality is maintained.

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11. Is the uptake of preservative solution by riceflower affected by the type of humectant used and its concentration? Introduction Riceflower (Ozothamnus diosmifolius syn. Helichrysum diosmifolium) is a member of the ‘paper daisy’ group of the Asteraceae family. It is a shrubby perennial species that produces masses of small flowers in attractive bunch-like inflorescences. These inflorescences are produced in spring. Whilst they are usually sold as a fresh cut flower, they can be easily air dried. There is, however, a loss of quality associated with air drying, as the material becomes very brittle. Thus it would be advantageous to develop appropriate treatments for the preservation of riceflower to extend the use and market of this Australian native cut flower crop. This chapter investigates the uptake of humectants by riceflower.

Objective 1. Investigate the effect of humectant type and concentration on uptake of solution by riceflower.

Methodology General methodology for investigating the rate of uptake, the amount of solution taken up and the relative change in mass Flowering stems of riceflower (Ozothamnus diosmifolius syn. Helichrysum diosmifolium) var. White Line 26 were harvested by Graham and Esther Cook from their property at Helidon. Uniform stems were selected, cut to 30 cm length and branches and leaves were removed from the lower 15 cm. All stems were weighed prior to the commencement of the experiment. Six humectant treatment solutions were used; 10%, 20% and 30% glycerol, 10% glycerol plus 10% polyethylene glycol (PEG), 15% glycerol plus 5% PEG and 20% PEG. All concentrations were determined on a volume to volume basis, and deionised water was used as a control. A biocide of 10mg/L available chlorine as dichloroisocyanurate, was added to each of the solutions. Ten replicates, each consisting of a cut stem in its own vase, were placed in a controlled environment at 24 ±1°C and 65 ±5% RH during experimentation. A completely randomised design was used. Solution uptake was recorded daily for 7 days unless otherwise stated. For each assessment, the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (the total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae: Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0

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where, t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time. The actual amount of glycerol taken up was calculated by multiplying the total weight of solution taken up by a factor based on the glycerol concentration and the specific gravity of glycerol (1.262). Thus for a 10% glycerol solution (v/v), the factor is 0.123. That is, it is equivalent to a 12.3% glycerol solution (w/w). For a 20% glycerol solution (v/v), the factor is 0.240, and 0.351 for a 30% solution (v/v). A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results pertaining to this chapter can be found in Appendix 11.

Detailed results This experiment showed that humectant type and concentration influence the uptake of solution by riceflower. In most cases, the uptake rate of the humectant solution was significantly slower that the control treatment of deionised water (Figure 11.1). There were two exceptions: at Day 1 the initial uptake rate for the 10% glycerol solution was significantly greater than the control (33.4 versus 26.7 mgg-1 fw h-1); and there was no significant difference between the control and the 20% glycerol solution. By Day 3, the uptake rate of the humectant solutions by the riceflower had declined to less than 10 mgg-1 fw h-1. The glycerol only solutions showed a trend of decreasing uptake rate with increasing glycerol concentration. Uptake of those solutions containing PEG was very slow. It is interesting to note that the addition of 10% PEG to a 10% glycerol solution halved the initial uptake rate compared with glycerol alone. There was a general increase in the cumulative uptake of solutions throughout the experimental period. The greatest amount of solution was taken up by those stems in the control treatment followed by the 10% glycerol treatment (Figure 11.2). All of the other treatments accumulated less than 1.2 gg-

1 fw. In considering the cumulative uptake by stems in the glycerol only solutions, there is once again a trend of decreasing uptake with increasing concentration of glycerol (Table 11.1). However, when the actual amount of glycerol taken up by stems is calculated, this trend is reversed. That is, significantly more glycerol was taken up by those stems in the more concentrated solutions. By Day 7, those stems in the 30% glycerol solution had accumulated 0.855 g of solution per gram of fresh weight, of which 0.300 g was glycerol, while those in the 10% solution had taken up twice as much solution (1.933 grams per gram of fresh weight) of which only 0.238 g was glycerol. After Day 5, there was no significant difference in the actual amount of glycerol per gram of fresh weight for stems in the 10% and 20% solutions. Table 11.1. Effect of solution glycerol concentration on cumulative uptake of glycerol compared with total solution uptake by riceflower Glycerol solution Uptake of: Day 1 Day 2 Day 3 Day 5 Day 7 10% Solution #0.800 1.359 1.555 1.759 1.933

Glycerol 0.099 0.167 0.191 0.216 0.238

20% Solution 0.5639 0.772 0.852 0.940 1.016

Glycerol 0.135 0.185 0.204 0.226 0.249

30% Solution 0.460 0.615 0.678 0.762 0.855

Glycerol 0.162 0.216 0.238 0.269 0.300

# Values are gg-1 fw

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The relative mass of the control treatment stems was initially very high, being 104% of initial weight. During the experiment, the relative mass of these stems declined to 68% at Day 7 (Figure 11.3). In contrast, those stems in the humectant treatments always had a relative mass of less than 100% of initial weight. The relative mass of these stems also declined, reaching a minimum around Day 3. In the later stages of the experiment, the relative mass of the humectant treated stems increased.

Figure 11.1: Effect over time of humectants on rate of solution uptake by riceflower (Ozothamnus diosmifolius) stems (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol; = 10% glycerol plus 10% PEG; x = 15% glycerol plus 5% PEG; o = 20% PEG). Bars show LSD (p=0.05) between means.

Figure 11.2: Effect of humectants on cumulative solution uptake by riceflower (Ozothamnus diosmifolius) stems (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol; = 10% glycerol plus 10% PEG; x = 15% glycerol plus 5% PEG; o = 20% PEG). Bars show LSD (p=0.05) between means.

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Figure 11.3: Effect over time of humectants on relative mass of riceflower (Ozothamnus diosmifolius) stems (υ = deionised water i.e. control; ν = 10% glycerol; σ = 20% glycerol; λ = 30% glycerol; = 10% glycerol plus 10% PEG; x = 15% glycerol plus 5% PEG; o = 20% PEG). Bars show LSD (p=0.05) between means. Discussion of results The presence of humectants in solution reduced the uptake of solution by stems of riceflower. This was particularly marked for the PEG treatments and is due to the large size of the PEG molecule. It was interesting to note the reversal in the trend associated with the uptake of glycerol solution in relation to glycerol concentration when the total solution uptake was compared with the uptake of the glycerol itself. Thus it would be possible to manipulate the concentration of the glycerol solution to fit a desired time frame for uptake of a predetermined amount of glycerol. This would allow processors to schedule the various procedures they undertake in order to achieve efficient and effective use of facilities and equipment. The results pertaining to the relative mass of the riceflower stems suggest that all stems underwent some wilting during the first half of the experimental period. Those stems in the control solution of deionised water were apparently unable to take up sufficient water during the second phase of the experiment to regain the loss in relative mass. In contrast those stems treated with the humectant solutions were able to recover to some degree. This may have been achieved by the humectants accumulated in the plant material absorbing water from what must have been an environment of higher relative humidity.

Implications For riceflower, uptake of humectant treatment must be completed within 3 days. For riceflower, it is possible to increase the concentration of the glycerol solution used and, thereby, reduce the time required to take up the required amount of glycerol. Polyethylene glycol does not seem to be a suitable humectant treatment for riceflower.

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Recommendations Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of riceflower. Tests also are required to identify if there is an upper limit to the concentration of the glycerol solution that can be used to satisfactorily preserve riceflower.

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12. What type of dye is most effective at colouring riceflower during uptake preservation? Introduction Plant materials preserved only with humectants tend to be dull in colour. Thus, dyes are often added to the preservative solution to enhance the attractiveness of the finished product. The choice of dye and the method of application are important considerations to ensure the desired effect is achieved. There are basically two different types of dyes: anionic and cationic. Anionic dyes, which include food dyes, dissolve readily in water due to their low molecular weight and negative charge (associated with SO3, COO- and O- ions). These dyes are frequently used for uptake colouring of plant material. In contrast, cationic dyes, such as the aniline or textile dyes, develop a positive charge in water because they contain basic groups (amines). These groups are relatively insoluble. Thus cationic dyes need to be applied at elevated temperatures to facilitate complete dissolution and are, therefore, applied to plant material using immersion techniques. Depending on the time, temperature and dye concentration, there is an infinite range of colours and intensities that can be achieved. There are three types of commercial dyes available for use by the flower-preserving industry: food dyes, textile dyes and ethanol-based Dutch dyes. This study aimed to determine which of these is most effective for colouring riceflower.

Objective 1. Evaluate the effectiveness of commercial dye types for colouring riceflower during uptake

preservation.

Methodology Flowering stems of riceflower (Ozothamnus diosmifolius syn. Helichrysum diosmifolium) var. Line 26 white were harvested from Graham and Esther Cook’s property at Helidon. Stems were returned to the laboratory and allowed to equilibrate to room temperature prior to experimentation. Red dyes were selected for each of the three dye types tested. Two solution types were used: 20% glycerol and deionised water, as a control. The food dye (Permicol Ponceau red, 5g/L) and the Dutch dye (enthanol-based red, 30ml/L) were added directly to the solutions and stirred in to dissolve. The food dye solution was then filtered to ensure that there were no particles left. The textile dye, Sandocryl red BRLN 200%, 5g/L, was first dissolved in hot water and filtered before addition to the glycerol solution. A biocide of 10mg/L available chlorine as dichloroisocyanurate was added to each of the solutions. Stems were re-cut under water to 30cm length. The leaves and branches were removed from the basal 15cm of stem and the stems were weighed immediately before placing in the treatment solutions. Ten replicates, each consisting of a cut stem in its own vase, were placed in a controlled environment at 24 ±1°C and 65 ±5% RH during experimentation. A completely randomised design was used. Solution uptake and stem weight were recorded daily for 8 days. The dye intensity in the plant material was noted. For each assessment, the solution uptake rate (the amount of solution taken up per hour), the cumulative uptake of solution (the total amount of solution taken up) and the relative mass of each stem were calculated using the following formulae:

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Weight of solution t=n-1 - Weight of solution t=n Uptake rate t=n (gg-1 fw h-1) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ ÷ n Weight of stem t=0 Weight of solution t=0 - Weight of solution t=n Cumulative uptake t=n (gg-1 fw) = ⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯⎯ Weight of stem t=0 Weight of stem t=n Relative mass t=n (% initial wt) = ⎯⎯⎯⎯⎯⎯⎯⎯ x 100 Weight of stem t=0 where t is the assessment time, n is the assessment time in hours and n-1 is the previous assessment time in hours. A factorial design was adopted to differentiate treatment means using the Balanced Model procedure for SYSTAT for Windows, Version 5 biometrics package (Evanston, 1992). Mean separation was by the least significant difference (LSD) test at p=0.05. A full set of statistical results for this chapter can be found in Appendix 12.

Detailed results There was a significant interaction between the solution i.e. 20% glycerol versus deionised water) and the type of dye used. Over all of the dye treatments the uptake rate and the accumulation of solution was significantly greater for the deionised water than for the glycerol solution. This tendency became more pronounced as the experiment proceeded over time (Figures 13.1 and 13.2). The addition of a dye significantly reduced the rate of solution uptake and accumulation in the plant material (Figure 12.1(a), 12.1(b), 12.2(a) and 12.2(b). There was no significant difference in uptake of the different dye treatments in the glycerol solution. However, the riceflower stems took up a greater and similar amount of solutions that contained either the Dutch dye or no dye at all, than those with the food and textile dyes, respectively. In contrast, for the dyes in the deionised water, significantly less solution was taken up in the presence of the food dye and significantly more in the absence of any dye. Differences in uptake between the Dutch and textile dyes were not significant. Observations at Day 3 showed that all of the stems in the deionised water dye solutions remained green. In contrast, stems in the glycerol-dye solutions were showing some discolouration. Those stems exposed to the food dye had turned red, as had the lower branches of stems in the textile dye. The stems in the solution containing the Dutch dye remained green, although the leaves had turned brown. Although not significantly different in the latter stages of the experiment, the relative mass of the glycerol treated stems was consistently lower than for stems in the deionised water treatments (Figures 12.3(a) and 12.3(b)). The relative mass of stems in deionised water declined throughout the experiment from an average of 102% to 84% of their initial weight. This effect was most marked in those stems exposed to the dyes. The relative mass of the stems in the glycerol solutions also declined in the initial stages of the experiment. By Day 5 these stems had recovered to about 84% of their initial weight and then remained at that level for the rest of the experimental period. In these stems there was no significant difference between the dye treatments.

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Figure 12.1(a): Effect over time of dye addition on rate of solution uptake by stems of riceflower (Ozothamnus diosmifolius) in deionised water (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye).

Figure 12.1(b): Effect over time of dye addition on rate of solution uptake by stems of riceflower (Ozothamnus diosmifolius) in 20% glycerol solution (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye). Bars show LSD (p=0.05) between means for significant interactions between dye treatment and solution.

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Figure 12.2(a): Effect of dye addition on cumulative solution uptake by stems of riceflower (Ozothamnus diosmifolius) in deionised water (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye).

Figure 12.2(b): Effect of dye addition on cumulative solution uptake by stems of riceflower (Ozothamnus diosmifolius) in 20% glycerol solution (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye). Bars show LSD (p=0.05) between means for significant interactions between dye treatment and solution.

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Figure 12.3(a): Effect over time of dye addition on relative mass of stems of riceflower (Ozothamnus diosmifolius) in deionised water (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye). Figure 12.3(b): Effect over time of dye addition on relative mass of stems of riceflower (Ozothamnus

diosmifolius) in 20% glycerol solution (υ = no dye i.e. control; ν = Dutch dye; σ = food dye; λ = textile dye). Bars show LSD (p=0.05) between means for significant interactions between dye treatment and solution.

Discussion of results In this study, the presence of the dyes did not significantly reduce the uptake of the glycerol preservative solution. Morover, there was no difference between the food, textile or Dutch dyes in

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terms of uptake. Despite this, those stems treated with the food dye showed far greater red staining of the leaf tissue than the other treatments. These were the only stems to show the colouration in the florets. The Ponceau food dye is particularly soluble in water because of its anionic nature and low molecular weight. These properties also allow it to be readily taken up by the plant material and distributed throughout the leaves and florets. In contrast, the textile dye and the Dutch dye did not dissolve as readily and may well have blocked the vascular tissue to some extent. These two dye types would be better used for immersion dying of plant material where it is possible to greatly increase the temperature of the solution and, thereby, improve the solubility and penetration of these chemicals It is interesting to note that the presence of the dyes also did not impact on the humecant properties of the glycerol solution. This was demonstrated by the recovery in relative mass across all of the glycerol treatments, irrespective of the presence or absence of the dye.

Implications Food dyes, added to uptake preservative solutions, may provide an appropriate, effective and efficient means of colouring the foliage and florets of riceflower. Textile dyes and Dutch dyes are inappropriate for use with uptake preservation solutions as a way of colouring the foliage and florets of riceflower, but may be effective during immersion dying.

Recommendations Further experimentation is required to determine if other coloured food dyes are also effective at colouring the foliage and florets of riceflower. Research is needed to determine the appropriate concentration and time of treatment to achieve a range of colour intensities.

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13. Conclusions and industry implications The participants of the initial industry workshops conducted in Queensland and Western Australia agreed that research was needed to address the key problems faced by the preserving industry in Australia. Hence this project was designed using a systematic scientific approach to identify appropriate chemicals and to optimise protocols for the uptake preservation of a selection of native cut flowers and foliages. The importance of fresh and healthy plant tissues for uptake preservation was shown. Stems to be preserved by uptake should be handled in a similar way to those stems to be sold as fresh cut flowers. The time between harvest and preservation needs to be minimised and stems should be recut before being placed into preservative solutions. Stems that have been harvested and handled incorrectly during transportation to the preservation shed are more likely to show variable or poor solution uptake. Species of Eucalyptus differ in their ability to take up water or glycerol solution. Thus, there may be a need to change the length of the preservative process on a per species basis. In one experiment under the same environmental conditions, E. tetragona required more time than E. robusta to take up the desired amount of solution and, therefore, glycerol. E. tetragona showed anatomical and morphological characteristics of adaptation to a xerophytic environment. However, physiological state of the stems at harvest and the prevailing environmental conditions may be more important than inherent morphological and anatomical characters, which may not necessarily affect the uptake of glycerol solution and hence preservation. The uptake of preservative solutions can be enhanced by modifying the prevailing environmental conditions during transportation, handling and processing. Appropriate conditions could result in more rapid and uniform solution uptake. Whilst most processors do not have elaborate environmentally controlled sheds in which to process cut stems of foliage, there are some very simple procedures and equipment which could be put in place to improve the outturn quality of stems preserved by uptake preservation and in turn its economic return. Stems of Eucalyptus foliage are best processed promptly, within 12 hours of harvest, and uptake of glycerol needs to be completed within 3 days. Prewilting of cut stems of E. robusta foliage for up to 12 hours prior to processing does not affect the uptake of glycerol. Thus, prewilting offers no advantage over processing material immediately after harvesting. Cut stems of E. robusta foliage can be encouraged to take up glycerol solution more rapidly by subjecting them to continuous elevated light intensities. Hence, elevated intensity lighting (e.g. 36 W Philips® cool white fluorescent lamps, to give 56-75 µEinm-2 s-1) should be installed approximately 25 cm above the foliage being processed. These lights should remain illuminated throughout the uptake process. Air circulation at low airspeeds can increase the uptake of glycerol solution by cut stems of E. robusta foliage. High airspeeds can be detrimental to the uptake of solution. Hence, fans should be installed in the processing area and operated on low during processing. The uptake of glycerol solution by cut stems of E. robusta foliage is improved by increasing the vapour pressure difference between the leaf and the atmosphere. This can be achieved by increasing the temperature during processing. Uptake is very poor at or below 7°C. Glycerol was found to be an appropriate preservative solution for E. robusta and E. tetragona. The most rapid accumulation of glycerol in stems of E. robusta and E. tetragona was achieved with a 20% glycerol solution. Uptake of glycerol solutions by E. robusta is reduced after 3 to 4 days, but this did not occur for E. tetragona. Polyethylene glycol (PEG), at the concentration and in the combination

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with glycerol tested, is an inappropriate humectant for the preservation of E. robusta and E. tetragona, as it caused discoloration and loss of flexibility of the foliage. Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of E. robusta and E. tetragona. The commercial adjuvants, Agrimul PG 2067 and Geropon SDS, at the concentration tested (100 mg mL-1), should not be added to a glycerol solution during uptake preservation of cut stems of either E. robusta or E. tetragona. The addition of Agrimul at this concentration does not provide any benefits and the addition of Geropon SDS was detrimental to the stems. Further experimentation is required to determine whether the use of adjuvants can provide any benefit to the uptake preservation process for E. robusta and E. tetragona. Such experimentation should consider the use of Agrimul PG 2067 and Geropon SDS at lower concentrations, as well as other adjuvants, like Agral 600. Testing needs to be done on a species by species basis. Pyranine dye can be used to assist in studying the movement of glycerol solution through cut stems of E. robusta. However, the degree of fluorescence caused by pyranine dye cannot be used as a measure of the amount of solution taken up by cut stems of E. robusta, unless it is standardised for each solution type. Based on the location of fluorescence in the stem and leaf tissues, glycerol moves relatively quickly and is distributed fairly evenly throughout these tissues over the initial 3 day period. In fact after as little as 24 hours of uptake of glycerol solution, the solution had been distributed to the peripheral leaves. Further experimentation is required to determine whether sufficient glycerol can be accumulated in all areas of the stem inside the 3 day period and still maintain the qualities expected of an uptake preservation solution in terms of foliage suppleness and colour. The plant growth regulators, ABA and fusicoccin, at the concentrations tested, should not be used to pre-treat cut stems of E. robusta for uptake preservation with glycerol solution. These plant growth regulators do not provide any benefits. Further experimentation is required to determine whether the use of these or other plant growth regulators may provide any benefit to the uptake preservation process for E. robusta. Such experimentation should consider varying the duration of the pulse, varying the concentration of the plant growth regulators in the pulsing or uptake solution, varying the application technique, (ie direct application to the foliage as a spray or as a dip), as well as using alternative chemicals that control stomatal aperture. Moreover, testing may need to be done on a species by species basis. Stenanthemum A 20% glycerol solution gave the most rapid accumulation of glycerol in stems of stenanthemum. However for stenanthemum, it is possible to increase the concentration of the glycerol solution used and thereby reduce the time required to take up the required amount of glycerol. Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of stenanthemum. Trials also are required to identify if there is an upper limit to the concentration of the glycerol solution that can be used to satisfactorily preserve stenanthemum. It is unclear whether polyethylene glycol alone or in combination with glycerol can be taken up by stenanthemum to a suitable level to provide any preservation of quality of the plant material. Further experimentation is required on fresh material of stenanthemum to determine whether polyethylene glycol alone or in combination with glycerol is a suitable humectant for preservation of this cut flower.

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Processors need to consider the environmental conditions during the preservation process and storage to ensure outturn quality of preserved stems is achieved and maintained. Storage of preserved flowering stems of at high relative humidity (>80%) and low relative humidity (30%) should be avoided as quality will be adversely affected by excessive water adsorption by humectants or by moisture loss respectively. If stems are to be packed for prolonged storage, cartons should have a low density polyethylene (LDPE) liner and a sachet of glucose or potassium chloride to maintain a stable environment and hence avoid loss of outturn quality. Adoption of the equilibrium relative humidity (ERH) test could determine the suitability of a product to the environmental conditions in the consumers display area. Alternatively it could be used as a feedback loop to refine the preservation process. Riceflower There are similarities in the uptake preservation behaviour of riceflower and stenanthemum, in that uptake of humectant treatment must be completed within 3 days. In addition, for both riceflower and stenanthemum, it is possible to reduce the time required to take up the required amount of glycerol by increasing the concentration of the glycerol solution used. Further experimentation is required to determine a recommended amount of glycerol per gram of fresh weight to achieve the desired preservation and quality of riceflower. Tests also are required to identify if there is an upper limit to the concentration of the glycerol solution that can be used to satisfactorily preserve riceflower. Polyethylene glycol does not seem to be a suitable humectant treatment for riceflower. Red food dyes, added to uptake preservative solutions, may provide an appropriate, effective and efficient means of colouring the foliage and florets of riceflower. Further experimentation is required to determine if other coloured food dyes are also effective at colouring the foliage and florets of riceflower. Research is needed to determine the appropriate concentration and time of treatment to achieve a range of colour intensities. Textile dyes and Dutch dyes are inappropriate for use with uptake preservation solutions as a way of colouring the foliage and florets of riceflower, but may be effective during immersion dyeing. The research conducted during this project has provide the dried and dye sectors of the Floricultural Industry with some useable results which can be incorporate simply into their own processing operations. Minor modifications to current practices can add to the efficiencies of the process, significantly improve outturn quality of these flowers and foliages and should ultimately result in better economic returns for the industry. The importance of using high quality products that were handled appropriately after harvest, as well as improving the environmental conditions during processing and understanding the different requirement for different species is of great importance.

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14. Bibliography Anon. (1996). Minitab Users Guide. Release 11.

(Minitab Inc: USA). Bhandari, B. (1997). Physio-chemical properties of

humectants. Proceedings of a workshop on preservation of cut flowers and foliage of Australian native plants. The University of Queensland Gatton College.

Campbell, S. (1996). ‘Preservation of Cut Eucalyptus

cinerea Foliage through Application of Glycerol’. Project (AG465) Bachelor of Horticultural Science, The University of Queensland, St Lucia.

Canny, M.J. (1995). Apoplastic water and solute

movement: new rules for an old space. Annual Review of Plant Physiology and Plant Molecular Biology. 46:215-236.

Dubois, P., and Joyce, D. (1990). Preserving plant

foliage with glycerine. (Western Australian Department of Agriculture: Perth, Australia).

Dubois, P., and Joyce, D. (1992). Preservation of

fresh cut ornamental plant material with glycerol. Postharvest Biology and Technology 2:145-153.

Esau, K. (1953). Plant Anatomy. (Wiley and Sons,

Inc.: New York, USA). Evanston, I.I. (Ed). (1992). SYSTAT for Windows:

statistics, version 5. SYSTAT Inc. Gartner, B.L. (1995). Plant Stem Physiology and

Functional Morphology. (Academic Press: New York, USA).

Hall, M.A. (1976). Plant Structure, Function and

Adaptation. (Macmillan Press Ltd.: London, UK).

Joyce, D. (1998). Dried and preserved ornamental

plant material: not new, but often overlooked and underrated. Acta Horticulturae 454:133-145.

Joyce, D. (1997). Equilibium relative humidity test applied to quality assurance for ornamental plant material preserved with humectants. Proceedings of a workshop on preservation of cut flowers and foliage of Australian native plants. The University of Queensland Gatton College.

Labuza, T.P (1984). ‘Moisture Sorption: Practical

Aspects of Isoterm Measurement and Use’. (American Association of Cereal Chemists: St Paul, Minnesota, USA).

Martin, J.T., and Juniper, B.E. (1970). The Cuticles

of Plants. (Edward Arnolds Publishers Ltd.: Edinburgh, UK).

Rockland, L.B. (1960). Saturated salt solutions for

static control of relative humidity between 5° and 40°C. Analytical Chemistry 32:1375-1376.

Salisbury, F.B., and Ross, C.W. (1992). Plant

Physiology. 4th Edition. (Wadsworth Publishing Co.: Belmont, USA).

Sebanek, J. (1992). Plant Physiology. (Elsevier

Science Pub. Co.: New York USA). Shirazi, A., and Cameron, A.C (1992). Controlling

relative humidity in modofied atmosphere packages of tomato fruit. HortScience 27(4):336-339.

Solomon, M.E. (1951). Control of humidity with

potassiun hydroxide, sulphuric acid, or other solutions. Bulletin of Entomological Research 42:543-559.

Taiz, L., and Zeiger, E. (1991). Plant Physiology.

(Benjamin/Cunnings Pub. Co. Inc.: California, USA).

Weast, R.C. (Editor-in-chief) (1977). ‘CRC

Handbook of Chemistry and Physics’. 58th Edition. (CRC Press: Boca Raton).

Willmer, C.M. (1983). Stomata. (Longman Inc.:

New York, USA). Winston, P.W., and Bates, D.H. (1960). Saturated

salt solutions for the control of humidity in biological research. Ecology 41(1): 232-237.

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15. Appendices

Appendix 1 Information that is available on preserving cut flowers and foliage and the factors affecting it

Introduction It is often difficult to know where to start looking for other information relating to a specific topic, in this case preserving cut flowers and foliage, or finding more detailed information to fully understand the processes involved. This chapter aims to provide a starting point for locating such information.

Objectives List reference material that may provide useful background information on: Processing methods for cut flowers and foliage (57 references) Chemicals used in preserving cut flowers and foliage (9 references) Plant water relations (19 references) Morphological characteristics of leaves and vascular tissue (21 references) Environmental factors that influence uptake preservation (15 references)

Reference lists Processing methods for cut flowers and foliage Anon. (1994). Blush on the rose. Florists’ Review.

185(8):55-86. Anon. (1996). R & D Plan: Wildflower & Native

Plant Program 1995-2000. (Rural Industries Research & Development Corporation: Canberra). 40pp.

Anon. (1996). Drying and processing techniques for

flowers. (University of Michigan: USA). Anon. (1996). Expanding drieds. Florists’ Review.

187 (8):120. Armitage, A.M. (1993). Drying and preserving. In

“Specialty cut flowers: the production of annuals, perennials, bulbs and woody plants for fresh and dried cut flowers”. (Varsity Press/Timber Press: Portland, USA).

Ashwell, M., and Pearson, S. (1995). Preserved and

artificial material. In “Professional Floristry Techniques”. (Farming Press: Ipswich, UK).

Barber, R. (1989). The delight of dried flowers. New

Zealand Gardener. 45 (2):10-11. Beveridge, A. (1996). The basics of freeze-drieds - the

process, the improvements and the limitations. Florists’ Review. 187(8):118.

Coates, B. (1995). Dried flower arrangements.

Australian Plants. 18 (144):143-150.

Dubois, P. (1990). Preserving plant foliage with glycerine. Farmnote No.87/90. Western Australian Department of Agriculture. 3pp.

Dubois, P. (1990). The use of biocides in glycerining

solutions. The Floriculture Industry Newsletter 13:17.

Dubois, P. (1990). Glycerining ornamental plant

foliage. The Floriculture Industry Newsletter 15:3.

Dubois, P., and Joyce, D. (1988). Bleaching plant

foliage. Farmnote No. 7/88. Western Australian Department of Agriculture. 3pp.

Dubois, P., and Joyce, D. (1989). Drying cut flowers

and foliage. Farmnote No. 10/89. Western Australian Department of Agriculture. 3pp.

Dubois, P., and Joyce, D. (1989). Dyeing cut flowers

and foliage. Farmnote No. 31/89. Western Australian Department of Agriculture. 4pp.

Dubois, P., and Joyce, D. (1992). Bleaching

ornamental plant material: a brief review. Australian Journal of Experimental Agriculture. 32:785-790.

Essex, J. (1986). All Cut and Dried. Hobbyfarmer.

October p. 38

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Fultz, C., and Bowser, D. (1985). Dry Ideas. Florists’ Review. November pp.16-22.

Given, P.S., Jr. (1991). Molecular behaviour of water

in a flour-water baked model system. In “Water Relationship in Foods: Advances in the 1980’s and Trends for the 1990’s”. (Levine, H., and Slade, L. Eds). (Plenum Press: New York). 863pp.

Gordon, L. and Lorimer, J. (Unknown). Preserving

and using unpressed flowers and foliage. “The Complete Guide to Drying and Preserving Flowers” pp.171-214. (Chartwell Books Inc.).

Greenway, H., and Leahy, M. (1970). Effects of

rapidly and slowly permeating osmotica on metabolism. Plant Physiology. 46:259-262.

Gulrajani, M.L., and Sukumar, N. (1985).

Optimisation of a single-stage preparatory process for cotton using sodium hypochlorite. Textile Research Journal. 56:614-619.

Gulrajani, M.L., and Venkatraj, R. (1986). A low

temperature scouring/ bleaching process for cotton using cotton chlorite. Textile Research Journal. 56:476-483.

Herschbach, D., and Stevens, A. (1999). Advances in

preserving decorative plant materials by the systemic absorption of glycerol and dye. New Flowers, Products and Technologies. Proceedings of the Fifth Australian Wildflower Conference. pp 86-90.

Heyser, J.W., and Nabors, M.W. (1981). Growth,

water content and solute accumulation of two tobacco cell lines cultured on sodium chloride, dextran and polyethylene glycol. Plant Physiology. 68:1454-1459.

Hillier, M. (1987). Malcolm Hillier’s Guide to

Arranging Dried Flowers. (Dorling Kindersley, Ltd: London). 160pp.

Hocking, O. (1995). Commercial Wildflower

Production “Dispelling the Myths”. Notes on Workshop Program. (Queensland Wax & Native Flower Association, and The University of Queensland, Gatton College). p.14.

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124

Appendix 2 Colour photographs of anatomical characteristics

Plate 15.2.1 A section of a leaf of Stenanthenum scortechinii showing trichomes on its undersurface, as well as two layers of elongated palisade mesophyll beneath the epidemis, spongy mesophyll, and a vascular bundle (Plate 2.1 in text)

Plate 15.2.2 Leaf anatomy of Eucalyptus tetragona showing palisade mesophyll on both sides of the leaf and a large oil gland on centre left of photo (Plate 2.2 in text).

125

Plate 15.2.3 Transverse section of Eucalptus sp. showing palisade mesophyll on upper side of the leaf. This was the arrangement for E. robusta, E. bakeri and E. crucis.

Plate 15.2.4 A photograph from scanning electron microscope image of Eucalyptus sp. leaf surface.

126

Plate 15.2.5 Transverse section of flowering stem of Stenanthemum scortechenii showing the many small xylem vessels (plate 2.2 in text)

Plate 15.2.6 Transverse section of Eucalyptus robusta showing xylem vessels that were typically larger than those for Stenanthemum scortechenii (Plate 9.2 in text).

127

Plate 15.2.7 Transverse section of Eucalyptus tetragona showing vessels of xylem and large oil glands in bark and pith.

128

Appendix 3 Statistical results pertaining to Chapter 2 Appendix table 15.3.1 Cumulative uptake of deionised water or glycerol, averaged over four Eucalyptus species. Solution Cumulative uptake (gg fw-1) Day 1 Day 2 Day 3 Day 4 Day 5 Day 7 Day 9 Deionised water

0.45 1.05

1.61

2.48

3.34

4.88

5.96

Glycerol

0.43

0.86

1.01

1.05

1.11

1.16

1.20

LSD0.05

NS

0.048

0.081

0.099

0.124

0.176

0.216

Appendix table 15.3.2 Cumulative solution uptake by four Eucalyptus species, averaged over solutions. Species Cumulative uptake of solution Day 1 Day 2 Day 3 Day 4 Day 5 Day 7 Day 9 E. robusta

0.67

1.24

1.61

2.05

2.50

3.33

3.88

E. tetragona

0.29

0..66

0.95

1.21

1.47

1.81

2.02

E. crucis

0.52

1.19

1.64

2.31

2.95

3.69

4.14

E. bakeri

0.27

0.71

1.03

1.50

1.97

3.24

4.29

LSD0.05

0.043

0.068

0.114

0.141

0.176

0.249

0.306