Host plant resistance as a management tool for Ditylenchus 
africanus (Nematoda: Tylenchidae) on groundnut (Arachis 
hypogaea) 
Sonia Steenkamp 
Thesis submitted for the degree Doctor of Philosophy at the Potchefstroom 
Campus of the North-West University 
South Africa 
Promoter: Prof AH Mc Donald 
Co-promoter: Prof D de Waele 
November 2008 
PREFACE 
Contributions, assistance and support by the following persons and institutions are 
gratefully acknowledged: 
My promoter Prof. Alex Mc Donald for his support and guidance. 
My co-promoter Prof. Dirk de Waele for his guidance and encouragement. 
Drs. Charlotte Mienie (ARC-GCI) and Liezel Herselman (University of the Free State) 
for their support and guidance in the molecular part of this study. 
Dr. Anine Jordaan (North West University) for her support and guidance in the 
histopathological part of this study. 
Ms Edith van den Berg (ARC biometrics division Pretoria) for her assistance in the 
statistical analysis of parts of this study. 
Prof. Koos van Rensburg (ARC-GCI) for his support, guidance, encouragement and 
help with all the stats. 
The ARC-Grain Crops Institute management for funding and support of this study. 
Dr. Driekie Fourie (ARC-GCI) for her inputs during this study. 
Erna Venter, Rita Jantjies, Nancy Ntidi, Belina Matule, Lizette Bronkhorst, Sameul 
Kwena and Abram Tladi for technical assistance. 
Dr. Piet van der Merwe, Dr. Willie Wentzel (ARC-GCI) and Sharmane Naido 
(University of KwaZulu Natal) for their patience in guiding me through the rougher 
parts of genetics. 
Mme. Alana Pretorius, Maria van der Merwe and Dr. Jan Dreyer of the groundnut 
breeding division (ARC-GCI) for their support, assistance and trust in this study. 
Meggie Kruger and Suria Bekker for collection and copying of literature. 
1 
Messrs. Schalk du Plessis and Derek Dean for the maintenance of the greenhouse. 
Mr. Peet van Heerden (PPECB) for providing infected groundnut pods from all over 
the groundnut producing area. 
Mr. Anton & Dick Howarth (Hartswater) and Willie Jansen (Jan Kempdorp) for 
providing field sites and assisting with the planting and maintenance of the trials. 
Wiltrud Durand for drawing of the map in Chapter 4. 
Mme. Mary James (public relations) and Petro Lombard (personal assistant) for 
helping with the printing of this thesis. 
My parents, family and friends for their support. 
11 
ABSTRACT 
HOST PLANT RESISTANCE AS A MANAGEMENT TOOL FOR DITYLENHUS 
AFRICANUS (MEMATODA: TYLENCHIDAE) ON GROUNDNUT {ARACHIS 
HYPOGAEA) 
Groundnut is an important cash crop both for commercial and smallscale farmers in 
South Africa. The effect of Ditylenchus africanus on groundnut is mainly qualitative, 
leading to downgrading of groundnut consignments. This nematode is difficult to 
control because of its high reproduction and damage potential. The objective of the 
study was to investigate the potential of host-plant resistance as an effective and 
economically-feasible alternative management tool for the control of D. africanus on 
groundnut. Selected groundnut genotypes were evaluated against D. africanus in 
microplot and field trials. PC254K1 and CG7 were identified as resistant to D. 
africanus. The resistance expressed by these two genotypes is sustainable under 
field conditions. The resistance expressed by PC254K1 is effective even at high 
population densities. This genotype consistently produced yields with a low UBS % at 
all nematode population levels. PC254K1 could therefore be used as a major source 
of resistance to D. africanus in the development of commercial cultivars. Although the 
breeding line PC287K5 also maintained low nematode numbers in some trials, its 
level of resistance does not seem to be as strong or as sustainable as that of 
PC254K1 or CG7. However, PC287K5 could still play an important role in the 
groundnut industry where lower D. africanus populations occur. The resistance 
expressed by PC254K1 is not transferred to leaf callus tissue of this genotype, 
confirming there is no short-cut for screening for resistance to D. africanus. The 
reproduction and damage potential of D. africanus populations from different 
geographically-isolated localities in the groundnut-production areas of South Africa 
was tested under controlled and semi-controlled conditions and were found to be 
similar to each other. Resistance of PC254K1 to all of the tested populations was 
confirmed. These results indicate that the presence of this resistant trait in a cultivar 
developed from PC254K1 should be sustainable over the whole groundnut-
production area of South Africa. The absence of D. africanus from pod tissue of 
PC254K1 confirmed the genotype's resistance. The mechanism of resistance 
involved may be the inhibition of proper development of this nematode, preventing it 
to build up to damaging population levels. However, PC254K1 is not immune to this 
nematode since it does occur in small numbers on this genotype. The resistance trait 
in PC254K1 is seemingly governed by a number of genes, implying that it will be 
in 
more durable under sustained pressure by D. africanus populations. Although 
markers associated with the resistance trait were mapped, they were not closely 
linked. Three putative qualitative trait loci (QTL's) were identified but markers 
associated with the resistance trait need to be refined and developed to be breeder-
friendly in terms of marker-assisted selection. There are strong indications that CG7, 
which is a parent of PC254K1, may have more superior levels of resistance to D. 
africanus than PC254K1. The identification of markers closely associated with the 
resistance trait might, therefore, be more successful using CG7 in stead of PC254K1. 
Key words: Arachis hypogaea, breeding, Ditylenchus africanus, groundnut, 
resistance, management. 
iv 
UITTREKSEL 
GASHEERPLANTWEERSTAND AS 'N BEHEERMATRIEEL VIR DITYLENCHUS 
AFRICANUS (NEMATODA: TYLENCHIDAE) OP GRONDBONE (ARACHIS 
HYPOGAEA) 
Grondbone is 'n belangrike kontantgewas beide vir kommersiele en kleinboere in 
Suid-Afrika. Die effek van Ditylenchus africanus op grondbone is hoofsaaklik 
kwalitatief en veroorsaak dat grondboonbesendings afgegradeer word. Hierdie 
aalwurm is moeilik om te beheer weens sy hoe voortplantings- en skadepotensiaal. 
Die doel van hierdie studie was om die potensiaal van gasheerplantweerstand as 'n 
effektiewe, ekonomies-aanvaarbare alternatiewe beheermaatrieel vir die beheer van 
D. africanus op grondbone te ondersoek. Geselekteerde grondboongenotipes is teen 
D. africanus ge-evalueer in mikroplot- en veldproewe. PC254K1 en CG7 is 
geidentifiseer om weerstandbiedend teen D. africanus te wees. Die 
weerstandseienskap van die twee genotipes is volhoubaar onder veldtoestande. 
Weerstand in PC254K1 is selfs effektief by hoe aalwurmbevolkingsvlakke en hierdie 
genotipe kan oeste met lae OGV % by alle bevolkingsvlakke produseer. PC254K1 
kan dus as 'n hoofbron van weerstand teen D. africanus gebruik word om nuwe, 
kommersiele kultivars te ontwikkel. Alhoewel die teellyn PC287K5 in sommige 
proewe lae aalwurmvlakke kon handhaaf, lyk dit asof laasgenoemde se vlak van 
weerstand nie so sterk of volhoudbaar is soos die van PC254K1 of CG7 nie. 
PC287K5 kan egter steeds 'n belangrike rol speel in die grondboonbedryf, veral in 
gebiede waar lae D. africanus besmettingsvlakke voorkom. Die weerstand wat in 
PC254K1 voorkom, word nie oorgedra na die blaarkallusweefsel nie, wat bevestig 
dat daar nie 'n kortpad is vir die evaluasie van weerstand teen D. africanus nie. Die 
voortplanting en skadepotensiaal van D. africanus bevolkings vanaf verskillende 
geografies-geisoleerde gebiede in die grondboon produksie area van Suid Afrika is 
onder beheerde en semi-beheerde toestande getoets. Die voortplanting en 
skadepotensiaal van die bevolkings was soortgelyk aan mekaar. Weerstand in 
PC254K1 is bevestig teen al die getoetste bevolkings wat daarop dui dat die 
weerstandbiedendheid van 'n kultivar wat uit PC254K1 geteel is, volhoubaar behoort 
te wees oor die hele grondboonproduksiegebied van Suid-Afrika. Die weerstand van 
PC254K1 word bevestig deur die afwesigheid van D. africanus in peulweefsel van 
die genotipe. Die meganisme van die weerstand is klaarblyklik dat die ontwikkeling 
van die aalwurm ge-inhibeer word en dus nie opbou na skadelike vlakke nie. 
PC254K1 is egter nie immuun teen die aalwurm nie omdat hierdie parasiete nog in 
v 
klein hoeveelhede in die saad van hierdie genotipe gevind kan word. Die 
weerstandbiedende eienskap van PC254K1 word skynbaar deur 'n aantal gene 
beheer. Dit impliseer dat die weerstand meer volhoubaar is onder konstante 
bevolkingsdruk deur D. africanus. Merkers wat met die weerstandseienskap 
geassosieer is, is gekarteer. Hierdie merkers is egter nie nou gekoppel met die 
eienskap nie. Drie tentatiewe veelvuldige eienskaplokusse (VEL) is ge'identifiseer. 
Merkers geassosieer met die weerstandseienskap moet egter verfyn en verder 
ontwikkel word in telervriendelike merkers, wat in merkerondersteunde seleksie 
gebruik kan word. Daar is sterk aanduidings dat CG7 moontlik superieure 
weerstandsvlakke bo die van PC254K1 mag he. Eersgenoemde is 'n ouer van die 
laasgenoemde en as CG7 in die plek van PC254K1 gebruik word, mag 
merkeridentifikasie dalk meer suksesvol uitgevoer word. 
Sleutelwoorde: Arachis hypogaea, beheer, Ditylenchus africanus, grondbone, teling, 
weerstand. 
VI 
CONTENTS 
CHAPTER 1 INTRODUCTION 1 
1.1 THE GROUNDNUT CROP
1.1.1 Origin and distribution of the crop 1 
1.1.2 Taxonomy of the crop 1 
1.1.3 Cultivation of the crop
1.1.4 Groundnut as a food source 2 
1.1.5 Production constraints
1.2 THE GROUNDNUT POD NEMATODE 4 
1.2.1 Origin and distribution
1.2.2 History 4 
1.2.3 Life cycle and reproductive potential 5 
1.2.4 Survival 5 
1.2.5 Symptoms and histopathology
1.2.6 Damage potential and economic importance 7 
1.3 NEMATODE MANAGEMENT STRATEGIES ON GROUNDNUT 9 
1.4 CURRENT NEMATODE MANAGEMENT TOOLS IMPLIMENTED 
FOR LOCAL GROUNDNUT PRODUCTION 10 
1.4.1 Nematicides 1
1.4.2 Cultural and biological management strategies 11 
1.4.3 Crop-based management strategies 1
1.4.3.1 Crop rotation 12 
1.4.3.2 Resistant cultivars
1.5 HOST PLANT RESPONSE 3 
1.5.1 Host sensitivity 1
1.5.2 Host efficiency
1.5.3 Genes expressing resistance 14 
1.6 MOLECULAR MARKERS 15 
1.7 RATIONALE AMD LAYOUT OF THE CURRENT STUDY 15 
vn 
CHAPTER 2 MATERIALS AND METHODS 18 
2.1 MICROPLOT FACILITIES 1
2.2 SOIL FOR GREENHOUSE AND MICROPLOT TRIALS 19 
2.2.1 Fumigation of soil 20 
2.2.2 Soil nutrients 2 
2.2.2.1 Greenhouse 2
2.2.2.2 Microplots
2.2.2.3 Field sites 3 
2.3 BIOLOGICAL MATERIALS 2
2.3.1 GERMPLASM
2.3.1.1 Seed 3 
2.3.1.2 Seed treatment 24 
2.3.2 NEMATODES 2
2.3.2.1 Nutrient medium and callus tissue culturing 25 
2.3.2.2 Inoculation of groundnut callus tissue with D. africanus 26 
2.3.2.3 Renewal of D. africanus callus tissue cultures 26 
2.3.2.4 Extraction of D. africanus from callus tissue 27 
2.3.2.5 Determination of the number of nematodes extracted 
from callus tissue 28 
2.3.2.6 Preparation of nematode inoculum and inoculation 
procedures
2.4 DATA COLLECTION AND TRIAL ASSESSMENT PROCEDURES 28 
2.4.1 Sampling 2
2.4.1.1 Greenhouse trials 28 
2.4.1.2 Microplot and field trials 29 
2.4.2 Nematode extractions 2
2.4.2.1 Soil samples 9 
2.4.2.2 Root and peg samples 30 
2.4.2.3 Hull and kernel samples 1 
2.4.3 Crop yield 3
2.4.3.1 Yield quantity 31 
2.4.3.2 Yield quality
2.5 STATISTICAL ANALYSES 3 
Vlll 
CHAPTER 3. IDENTIFICATION OF GROUNDNUT GENOTYPES WITH 
RESISTANCE TO DITYLENCHUS AFRICANUS 34 
3.1 INTRODUCTION 3
3.2 MATERIALS AND METHODS 35 
3.2.1 Identification of D. a/r/canus-resistant groundnut genotypes 35 
3.2.1.1 Trial layout 6 
3.2.1.1.1 Seed and genotypes 3
3.2.1.1.2 Microplot trials 37 
3.2.1.1.3 Field trials 8 
3.2.1.2 Collection of nematode and yield data 39 
3.2.2 Reproduction and damage-threshold levels of D. africanus 
on resistant, susceptible and tolerant groundnut genotypes 40 
3.2.3 Expression of D. africanus resistance in callus tissue 41 
3.3 STATISTICAL ANALYSES 42 
3.4 RESULTS 43 
3.4.1 Identification of D. a/r/canus-resistant groundnut genotypes 43 
3.4.1.1 Microplot trials 4
3.4.1.1.1 Final nematode population densities, 
reproduction factor and reproduction rate 43 
3.4.1.1.2 Yield assessments 46 
3.4.1.2 Field trials 50 
3.4.1.2.1 Final nematode population densities, 
reproduction factor and reproduction rate 50 
3.4.1.2.2 Yield assessments 53 
3.4.2 Reproduction and damage-threshold levels of D. africanus 
on resistant, susceptible and tolerant groundnut genotypes 54 
3.4.2.1 Final nematode population densities and reproduction 
factor 5
3.4.2.2 Yield assessments 56 
3.4.3 Expression of D. africanus resistance in callus tissue 59 
3.4.3.1 Final nematode population densities and reproduction 
factor 5
3.5 DISCUSSION 60 
IX 
CHAPTER 4. REPRODUCTIVE AND DAMAGE POTENTIAL OF FIVE 
GEOGRAPHICALLY-ISOLATED DITYLENCHUS AFRICANUS POPULATIONS ON 
GROUNDNUT 64 
4.1 INTRODUCTION
4.2 MATERIALS AND METHODS 65 
4.2.1 D. africanus populations
4.2.2 Trial layout 66 
4.2.2.1 Growth cabinet trial 6
4.2.2.2 Greenhouse trial 7 
4.2.2.3 Microplot trial 6
4.3 STATISTICAL ANALYSES 8 
4.4 RESULTS 69 
4.4.1 Growth cabinet trial 6
4.4.1.1 Final nematode population densities and reproduction 
factor 9 
4.4.2 Greenhouse trial 71 
4.4.2.1 Final nematode population densities and reproduction 
factor
4.4.3 Microplot trial 73 
4.4.3.1 Final nematode population densities and reproduction 
factor
4.4.3.3 Damage assessments 74 
4.5 DISCUSSION 75 
CHAPTERS. THE MECHANISM OF RESISTANCE TO DITYLENCHUS 
AFRICANUS EXPRESSED BY THE GROUNDNUT GENOTYPE PC254K1 78 
5.1 INTRODUCTION 7
5.2 MATERIALS AMD METHODS 78 
5.2.1 Trial layout in the greenhouse 7
5.2.2 Preparation of plant tissue for light-microscope studies 79 
5.3 RESULTS 80 
5.3.1 External damage symptoms on mature pods collected at 150 DAP 80 
5.3.2 Histopathology 81 
5.3.2.1 Pegs
5.3.2.2 Pods 4 
x 
5.4 DISCUSSION 89 
CHAPTER 6. ORIGIN OF RESISTANCE TO DITYLENCHUS AFRICANUS IN 
PC254K1 AND IDENTIFICATION OF MOLECULAR MARKERS 91 
6.1 INTRODUCTION 9
6.2 MATERIALS AND METHODS 92 
6.2.1 Origin of the resistance trait 9
6.2.1.1 First-generation crosses (FT seed) 92 
6.2.1.2 Second-generation (F2) seed 94 
6.2.1.3 Evaluation of the F2 progeny for D. africanus resistance 96 
6.2.2 Molecular marker identification 97 
6.2.2.1 DNA extraction
6.2.2.2 Amplified fragment length polymorphism (AFLP) analysis 97 
6.2.2.3 Bulk segregant analysis 99 
6.3 STATISTICAL ANALYSES 100 
6.3.1 Origin of the resistant trait 10
6.3.2 Marker identification 10
6.4 RESULTS 101 
6.4.1 Origin of the resistant trait 10
6.4.2 Molecular marker identification 104 
6.4.2.1 AFLP analysis 10
6.4.2.2 Linkage analysis 4 
6.4.2.3 QTL analysis 108 
6.5 DISCUSSION 10
CHAPTER 7. CONCLUSIONS 110 
CHAPTER 8. BIBLIOGRAPHY 113 
XI 
LIST OF FIGURES 
Figure 1.1. Initial symptoms on groundnut pods caused by Ditylenchus africanus 
infection appears at the connection of the peg to the pod. 6 
Figure 1.2. Ditylenchus africanus on the cotyledon and embryo of an infected 
groundnut kernel. 7 
Figure 1.3. Groundnut plants are lifted at harvesting. In severely-infested 
Ditylenchus africanus fields, 40 to 60% of the pods breaks off and 
remains behind in the soil. 8 
Figure 1.4. Groundnut seed and pods infected with Ditylenchus africanus. 8 
Figure 2.1. A micoplot facility of 20 rectangular brick troughs covered with a hail 
net. 18 
Figure 2.2. A microplot facility of evenly-spaced, concrete pipes and hail-net 
cover. 19 
Figure 2.3. A special hand applicator used for EDB fumigation of soil. 20 
Figure 2.4. Operators wearing full protective clothing during fumigation of soil 
with EDB in the microplots. 21 
Figure 2.5. Nutrients are incorporated into the top 30 cm of the soil with a 
garden fork. 22 
Figure 2.6. Application of herbicides with a tractor-mounted implement. 23 
Figure 2.7. A. Ditylenchus africanus groundnut callus tissue cultures such as 
those used for the inoculation of microplot and greenhouse trials in 
this study. B. Growth cabinets used for the incubation of 
Ditylenchus africanus groundnut callus tissue cultures. 25 
Figure 2.8. A. A mature four-month-old groundnut callus. B. A Ditylenchus 
africanusAnfested groundnut callus tissue culture due for renewal. 27 
xn 
Figure 3.1. A groundnut field trial at Jan Kempdorp. 38 
Figure 3.2. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from the inoculated section of a microplot trial during 
2003-2004 (P < 0.05; F-ratio = 10.02). 44 
Figure 3.3. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from the inoculated section of a microplot trial during 
2004-2005 (P < 0.05; F-ratio = 40.69). 44 
Figure 3.4. Ditylenchus africanus numbers (Pf) in the roots of eight groundnut 
genotypes from a field trial planted at Hartswater during 
2004-2005 (P < 0.05; F-ratio = 4.43). 50 
Figure 3.5. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from a field trial planted at Hartswater during 
2004-2005 (P < 0.05; F-ratio = 15.38). 51 
Figure 3.6. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from a field trial planted at Jan Kempdorp during 
2004-2005 (P < 0.05; F-ratio = 44.06). 52 
Figure 3.7. Non-linear relationships between initial nematode population 
densities (Pi) and final nematode population densities (Pf) in 
pods of Sellie, Kwarts, PC254K1 and PC287K5 in a microplot 
trial during 2004-2005 56 
Figure 3.8. Relationships between initial nematode population densities (Pi) 
and UBS % of Sellie, Kwarts, PC254K1 and PC287K5 in a 
microplot trial during 2004-2005 58 
Figure 4.1. Ditylenchus africanus populations were isolated from infected 
groundnut pods collected from five different localities (indicated by 
red dots) within the groundnut-production area of South Africa. 65 
xm 
Figure 4.2. Final D. africanus numbers (Pf) on groundnut callus tissue 
inoculated with five geographically-isolated Ditylenchus 
africanus populations from Mareetsane (population 1), Jan 
Kempdorp (population 2), Vaalharts (population 3), 
Schweizer-Renecke (population 4) and Theunissen (population 5) 
incubated for four weeks at 21 °C, 28 °C or 35 °C in three 
separate growth cabinets during 2007 (P < 0.05; F-ratio = 4.21). 69 
Figure 4.3. Final D. africanus numbers (Pf) on groundnut callus tissue 
inoculated with five geographically-isolated Ditylenchus 
africanus populations from Mareetsane (population 1), Jan 
Kempdorp (population 2), Vaalharts (population 3), 
Schweizer-Renecke (population 4) and Theunissen (population 5) 
pooled over three temperature regimes, respectively, in a growth 
cabinet trial during 2007 (P < 0.05; F-ratio = 76.69). 70 
Figure 4.4. Final Ditylenchus africanus numbers (Pf) in pods of Sellie 
and PC254K1 inoculated at planting with five geographically-
isolated Ditylenchus africanus populations from Mareetsane 
(population 1), Jan Kempdorp (population 2), Vaalharts 
(population 3), Schweizer-Renecke (population 4) and 
Theunissen (population 5) in a greenhouse trial during 2007 
(P < 0.05; F-ratio = 2.92). 72 
Figure 4.5. Final Ditylenchus africanus numbers (Pf) in pods of Sellie 
and PC254K1 inoculated at planting with five geographically-
isolated Ditylenchus africanus populations from Mareetsane 
(population 1), Jan Kempdorp (population 2), Vaalharts 
(population 3), Schweizer-Renecke (population 4) and 
Theunissen (population 5) in a microplot trial during 2007-2008 
(P < 0.05; F-ratio = 75.76). 73 
Figure 5.1. External symptoms on Sellie and PC254K1 pods 150 DAP at 
an inoculum level of 7 000 Ditylenchus africanus specimens 
in a greenhouse trial during 2008. 81 
xiv 
Figure 5.2. Light micrograph of a longitudinal section of a peg from A. Sellie 
and B. PC254K1 at inoculum level of 7 000 Ditylenchus africanus 
specimens at 150 DAP from a histolopathology study during 2008. 
A & B magnified 20 x. D = developing fibre layer, EN = endocarp, 
ES = embryo sac, EX = exocarp, F = fruit locule, M = mesocarp, 
O = ovule, V = vascular bundle 82 
Figure 5.3. Light micrograph of a cross section through the vascular region of 
pegs from A. Sellie and B. PC254K1 at inoculum level of 7 000 
Ditylenchus africanus specimens at 150 DAP from a histopathology 
study during 2008. A & B magnified 100 x. DUM = damaged 
undifferentiated tracheary elements of the metaxylem, 
DP = differentiated tracheary elements of the protoxylem, 
EN = endocarp, M = mesocarp, UM = undifferentiated tracheary 
elements of the metaxylem. 83 
Figure 5.4. Light micrograph of a cross section through A. The vascular bundle 
of the funicle and surrounding endocarp and mesocarp and B. The 
vascular bundle of the funicle and surrounding endocarp of a mature 
Sellie pod at inoculum level of 7 000 Ditylenchus africanus specimens 
at 150 DAP from a histopathology study during 2008. A magnified 
20 x and B 40 x. EN = endocarp, EX = eksocarp, FM = fibrous 
mesocarp, M = mesocarp, N = nematode, V = vascular bundle of the 
funicle. 85 
Figure 5.5. Light micrograph of a cross section through A. The vascular region 
and surrounding mesocarp and B. The vascular region of a mature 
PC254K1 pod at inoculum level of 7 000 Ditylenchus africanus 
specimens at 150 DAP from a histopathology study during 2008. 
A magnified 20 x, B 40 x. EN = endocarp, FM = fibrous mesocarp, 
M = mesocarp, N = nematode, V = vascular bundles. 87 
xv 
Figure 5.6. Light micrograph of a cross section through the cells of the vascular 
region of a mature PC254K1 pod at inoculum level of 7 000 
Ditylenchus africanus specimens at 150 DAP from a histopathology 
study during 2008. Magnified 100 x. DUM = damaged 
undifferentiated tracheary elements of the metaxylem, 
DP = differentiated tracheary elements of the protoxylem, 
M = mesocarp, N = nematode. 88 
Figure 6.1. Emasculated flowers on a groundnut plant marked with coloured 
strings for artificial pollination. 93 
Figure 6.2. Female plants covered with a plastic bag supported by a wire frame. 94 
Figure 6.3. Groundnut breeding facility in a greenhouse. 95 
Figure 6.4. Mature pods produced by Fi plants after crosses were made 
between Sellie and PC254K1. 95 
Figure 6.5. Distribution of the F2 plants from a cross between susceptible Sellie 
($) and resistant PC254K1 (<$) according to the mean number of 
Ditylenchus africanus ln(x+1) (Pf) in pods per plant at harvesting of 
a microplot trial during 2006-2007. 103 
Figure 6.6. Distribution of the F2 plants from a cross between resistant PC254K1 
(S) and susceptible Sellie ($) according to the mean number of 
Ditylenchus africanus ln(x+1) (Pf) in pods per plant at harvesting of a 
microplot trial during 2006-2007. 103 
Figure 6.7. AFLP-based, genetic-linkage map developed for groundnut using a 
F2 population resulting from a cross between the Ditylenchus 
africanus- resistant PC254K1 and susceptible Sellie. Markers are 
indicated on the right of each bar and the distances between each 
marker are indicated in centi Morgan (cM) on the left of each bar. 105 
xvi 
Figure 6.8. QTL-likelihood plot of markers for Ditylenchus africanus resistance 
on linkage groups (LG) 4 & 6. Markers are indicated to the right of 
each LG bar and the distances between each marker in centi Morgan 
(cM) to the left of the bar. Likelihood plots generated by 
MAPMAKER.QTL are indicated in blocks. The putative QTLs 
(D. africanus 1 & 2) are indicated by solid black bars. 108 
xvn 
LIST OF TABLES 
Table 2.1. Local specifications for groundnut grading as stipulated by the Act 
on Agricultural Product Standards, 119 of 1990 (SA, 1997). 32 
Table 3.1. Origin and preferred characteristics of groundnut genotypes that 
were selected for the identification of resistance to Ditylenchus 
africanus. 36 
Table 3.2. Groundnut entries for the identification of resistance to Ditylenchus 
africanus in the respective mircoplot and field trials conducted over 
two consecutive growing seasons (2003-2004 - 2004-2005). 37 
Table 3.3. Ditylenchus africanus numbers (Pf), reproduction factors (RF) and 
reproduction rates (RR) in pods of eight groundnut genotypes from 
a microplot trial during 2003-2004. 45 
Table 3.4. Ditylenchus africanus numbers (Pf), reproduction factors (RF) 
and reproduction rates (RR) in pods of eight groundnut genotypes 
from a microplot trial during 2004-2005. 46 
Table 3.5. Yield quality of eight groundnut genotypes in a microplot trial during 
2003-2004. 47 
Table 3.6. Yield quality of eight groundnut genotypes in a microplot trial during 
2004-2005. 47 
Table 3.7. Yield and income loss or gain of eight groundnut genotypes from 
inoculated and uninoculated sections of a microplot trial during 
2003-2004. 48 
Table 3.8. Yield and income loss or gain of eight groundnut genotypes from 
inoculated and uninoculated sections of a microplot trial during 
2004-2005. 49 
xvin 
Table 3.9. Ditylenchus africanus numbers (Pf) and reproduction rates (RR) in 
pods of eight groundnut genotypes from field trials planted at 
Hartswater and Jan Kempdorp during 2004-2005. 52 
Table 3.10. Yield quality of eight groundnut genotypes from a field trial planted 
at Hartswater during 2004-2005. 53 
Table 3.11. Yield quality of eight groundnut genotypes from a field trial planted 
at Jan Kempdorp during 2004-2005. 54 
Table 3.12. Final population densities (Pf) and reproduction factor (RF) of 
Ditylenchus africanus in pods of four groundnut genotypes 
inoculated at planting with escalating initial population densities (Pi) 
in a microplot trial during 2004-2005. 55 
Table 3.13. Yield quality and income for four groundnut genotypes inoculated at 
planting with a range of initial Ditylenchus africanus population 
densities (Pi) in a microplot trial during 2004-2005. 57 
Table 3.14. Yield of four groundnut genotypes inoculated at planting with a 
range of initial Ditylenchus africanus population densities (Pi) in 
a microplot trial during 2004-2005. 59 
Table 3.15. Ditylenchus africanus numbers (Pf) and reproduction factors (RF) 
on callus tissue of four groundnut genotypes incubated four weeks 
at 28 °C in a growth cabinet trial during 2008. 60 
Table 4.1. Reproduction factor (RF) of five geographically-isolated Ditylenchus 
africanus populations from Mareetsane (population 1), Jan Kempdorp 
(population 2), Vaalharts (population 3), Schweizer-Renecke 
(population 4) and Theunissen (population 5) on groundnut callus 
incubated four weeks at 21 °C, 28 °C and 35 °C in separate growth 
cabinets during 2007. 71 
xix 
Table 4.2. Reproduction factors (Pf) of five geographically-isolated Ditylenchus 
africanus populations from Mareetsane (population 1), Jan Kempdorp 
(population 2), Vaalharts (population 3), Schweizer-Renecke 
(population 4) and Theunissen (population 5) in pods of Sellie 
and PC254K1 in a greenhouse trial during 2007. 72 
Table 4.3. Reproduction factor (Pf) of five geographically-isolated Ditylenchus 
africanus populations from Mareetsane (population 1), Jan Kempdorp 
(population 2), Vaalharts (population 3), Schweizer-Renecke 
(population 4) and Theunissen (population 5) in pods of Sellie and 
PC254K1 in a microplot trial during 2007-2008. 74 
Table 4.4. Yield quality of Sellie and PC254K1 inoculated at planting with five 
geographically-isolated Ditylenchus africanus populations from 
Mareetsane (population 1), Jan Kempdorp (population 2), Vaalharts 
(population 3), Schweizer-Renecke (population 4) and Theunissen 
(population 5) in a microplot trial during 2007-2008. 75 
Table 6.1. Mlu\ (Ml) and Mse\ (M) adapter, primer+1 and primer+3 sequences 
used in amplified fragment length polymorphism (AFLP) analyses 
of genomic deoxyribonucleic acid (DNA) of PC254K1, Sellie and the 
F2 generation resulting from a PC254K1 x Sellie cross. 98 
Table 6.2. Percentage reproduction rate (RR) of Ditylenchus africanus on pods 
of F2 plants from Sellie ($) x PC254K1 (3) cross and its reciprocal 
PC254K1($) x Sellie (<$) from a microplot trial during 2006-2007. 102 
Table 6.3. AFLP primers tested on the DNA of the parents PC254K1 (resistant) 
($) and Sellie (susceptible) (c?) and the F2 progeny from PC254K1 x 
Sellie. 104 
Table 6.4. Linear regression analysis of DNA markers on the seven linkage 
groups (LG) on the AFLP-based genetic linkage map of groundnut. 
Loci were ordered using MAPMAKER.EXP and positions for putative 
QTLs for Ditylenchus africanus resistance were determined using 
MAPMAKER.QTL 107 
xx 
CHAPTER 1 
INTRODUCTION 
1.1 THE GROUNDNUT CROP 
1.1.1 Origin and distribution of the crop 
The genus Arachis is native to South America and probably originated in central 
Brazil (Stalker & Moss, 1987) or northeast Paraguay (Simpson et al., 2001). Wild 
Arachis species are distributed in a relatively small area that stretches between the 
Atlantic Ocean and the foothills of the Andes, from the mouth of the Amazon in the 
north to Uruguay in the south (Stalker & Moss, 1987). Gregory et al. (1973) 
suggested that most ancient Arachis species were found at high elevations. 
However, the geocarpic habit of Arachis suggests that long-distance dispersal 
occurred through water courses (Stalker & Moss, 1987) and that more recent 
speciation occurred as seeds were washed towards the sea (Gregory et al., 1973). 
As the seeds dispersed to lower elevations they became isolated in major river 
valleys and different sections of the genus evolved in parallel evolution (Gregory et 
al., 1973). The cultivated species A. hypogaea L. probably originated from the wild 
allotetraploid species A. monticola Krap. et Rig. since the latter is the only tetraploid 
known to be cross-compatible with A. hypogaea (Stalker & Moss, 1987). 
1.1.2 Taxonomy of the crop 
Arachis species fit into nine taxonomic sections including the cultivated groundnut A. 
hypogaea, which is subdivided into the subspecies hypogaea and fastigiata 
(Krapovicas & Gregory, 1994). Sub-species hypogaea includes the botanical 
varieties (var.) hypogaea (Virginia) and var. hirsuta (Peruvian runner) and subspecies 
fastigiata var. fastigiata (Valencia) and var. vulgaris (Spanish type) (Knauft & Wynne, 
1995). 
1.1.3 Cultivation of the crop 
A. hypogaea is a self-pollinating, annual, herbaceous legume (Hammons, 1982) and 
is the only Arachis species cultivated extensively for commercial production of seed 
and oil (Stalker & Moss, 1987). A. hypogaea flowers aboveground but carries its 
1 
pods beneath the soil surface in which two to four kernels are formed per pod, 
depending on the variety and cultivar (Dickson & De Waele, 2005). Groundnut is 
cultivated over six continents (Dickson & De Waele, 2005) in a wide range of field 
conditions ranging from clays to sands and from acidic to alkaline soils (Stalker & 
Moss, 1987). Globally, approximately 20 million hectares are under groundnut 
cultivation (Carley & Fletcher, 1995) in approximately 100 countries with tropical, 
sub-tropical and warmer temperate climates (Naidu et ai, 1999). 
Seventy two percent of the world's groundnut supply is produced in the Peoples' 
Republic of China, India, USA, Indonesia, Argentina, Senegal, Zaire and Myanmar 
(Dickson & De Waele, 2005). Global production averages nearly 24 million metric 
tonnes (Carley & Fletcher, 1995) and a significant proportion thereof is grown by 
resource-poor, smallholder farmers in developing countries (McDonald et ai, 1998). 
In Africa, approximately 5.3 million hectares are cultivated with groundnut (Carley & 
Fletcher, 1995) and major producers include Nigeria, Senegal, South Africa, Sudan, 
Zaire and the Democratic Republic of Congo (De Waele & Swanevelder, 2001). 
1.1.4 Groundnut as a food source 
Groundnut is listed as one of 20 crops standing between man and starvation 
(Wittwer, 1981). The calorie-rich kernels contain 25 % protein, 50 % oil, 20 % 
carbohydrate and 5 % fibre and ash (Knauft & Wynne, 1995) and may be boiled, 
broiled, roasted, fried, ground into butter, used as confectionary or crushed for oil 
(Dickson & De Waele, 2005). In the largest part of sub-Saharan Africa groundnut is 
an important subsistence crop grown mostly under rain-fed conditions (Van der 
Merwe et ai, 2001). Groundnut is primarily used as a cash crop and even 
smallholder farmers may sell their entire harvest (Stalker & Moss, 1987), which 
contributes significantly to food security and the alleviation of poverty in some 
countries and communities (Smartt, 1994). In South Africa groundnut is an important 
cash crop both for commercial and smallholder farmers (Mc Donald et ai, 2005). 
1.1.5 Production constraints 
Since groundnut is either a processed or a directly consumable food source, 
optimum kernel quality is important at all levels of production and utilisation (Hinds et 
ai, 1992; Swanevelder, 1997). Groundnut production and kernel quality can be 
limited by many abiotic and biotic factors, however. Drought is the most critical 
2 
abiotic factor that limits yield (Stalker, 1997). Yield constraints can also be caused by 
a calcium deficiency (Sumner et al., 1988), phosphorus-deficient soils and a less-
than-optimal relationship between nitrogen-fixing bacteria and the plant, inefficient 
nitrogen fixation due to environmental conditions (Knauft & Wynne, 1995), herbicidal 
and other chemical injuries or nutrient imbalances (Heagle, 1997). Iron, zinc, 
magnesium or boron deficiencies that occur in localised areas with a high pH also 
place constraints on groundnut production (Stalker, 1997; De Waele & Swanevelder, 
2001). Biotic factors affecting yields include weeds (Knauft & Wynne, 1995), insects 
(Isleib et al., 1994; De Waele & Swanevelder, 2001), diseases (Knauft et a/., 1988; 
Sharma & McDonald, 1990; Subrahmanyam etal., 1990; Reddy, 1991; Mehan et al., 
1995; Murant et a/., 1995; McDonald et a/., 1998; De Waele & Swanevleder, 2001) 
and nematodes (Stalker & Moss, 1987; Kokalis-Burelle et al., 1997; Dickson & De 
Waele, 2005). 
Worldwide, plant-parasitic nematodes are primary parasites of groundnut (Dickson & 
De Waele, 2005) and are able to cause detrimental losses in groundnut production 
(Kokalis-Burelle et al., 1997). New nematode species that cause crop damage 
continue to be discovered worldwide (Barker et al., 1994) and may also be 
associated with high levels of aflatoxin and other soil-borne diseases (Porter et al., 
1982; Timper et al., 2003). Annual losses caused by nematodes are globally 
estimated at 10 to 12 % when various crops are considered (Kahn, 2008), which 
translates into monetary losses of approximately US$6 billion in the USA alone 
(Agrios, 2005). However, the impact of plant-parasitic nematodes may be even 
higher than estimated since plant symptoms of nematode damage are usually 
nonspecific and yield losses caused by plant-parasitic nematodes often go unnoticed 
(Barker et al., 1994). 
Economic losses caused by nematodes will be much higher without the application of 
various nematode-management strategies and tactics (Barker et al., 1994). In many 
regions of the world groundnut cannot be grown without the effective management of 
nematode populations (Porter et al., 1982). Although many plant-parasitic nematodes 
have been associated with groundnut production locally (Venter et al., 1992) 
nematodes were not considered to be a major pest until the discovery of the 
groundnut pod nematode, Ditylenchus africanus Wendt, Swart, Vrain and Webster, 
1995 (Jones & De Waele, 1988; De Waele et al., 1989). 
3 
1.2 THE GROUNDNUT POD NEMATODE 
1.2.1 Origin and distribution 
D. africanus was first discovered on severely damaged hulls and seeds of groundnut 
collected from a rain-fed field in the Schweizer-Renecke district (27.19°S, 25.33°E), 
North West Province, South Africa during May 1987 (Jones & De Waele, 1988; De 
Waele et al., 1989). A national survey conducted during 1989 implies that this 
groundnut-parasitising nematode was present in the whole groundnut-producing area 
(De Waele et al., 1989). Of the 877 seed samples collected during this survey that 
were graded as damaged, 73 % was infected with this nematode (De Waele et al., 
1989). 
1.2.2 History 
D. africanus was originally identified as D. destructor Thome 1945 (Jones & De 
Waele, 1988), which is an important pest of potato tubers and flower bulbs in 
temperate regions of Europe, the USSR and localised areas in the USA (Hooper & 
Southey, 1982). A molecular study on the comparative taxonomy between some 
Ditylenchus populations (Wendt, 1992) and analysis of the ribosomal DNA (rDNA) of 
several geographic and host isolates of D. dipsachi Filipjev, 1936, D. myceliophagus 
Goodey, 1958 and D. destructor (Wendt et al., 1993) however, casted doubt on the 
original classification. The local Ditylenchus associated with groundnut, furthermore, 
did not damage potato tubers (De Waele et al., 1991) or other crops (De Waele et al., 
1989) and was consequently considered to belong to a different ecotype (De Waele 
& Wilken, 1990) that formed a distinct D. destructor race with a limited host range 
(De Waele et al., 1991). Wendt and Webster (1992) also indicated that the rDNA of 
D. destructor specimens from South Africa differed from that of D. destructor 
specimens from the United Kingdom and Wisconsin, USA. Based on the 
characteristics of morphology and restriction fragment length polymorphisms 
(RFLP's) of ribosomal DNA (rDNA), D. africanus was finally described in 1995 as a 
new Ditylenchus species (Wendt et al., 1995) that parasitizes various crops (Basson 
et al., 1990) and weeds (De Waele et al., 1990 & 1997) but causes damage only to 
groundnut (De Waele et al., 1989). So far D. africanus has not yet been reported on 
groundnut from other parts of the world and it seems to be endemic to South Africa 
(Dickson & De Waele, 2005). 
4 
1.2.3 Life cycle and reproductive potential 
Most of D. africanus' eggs (up to 13 eggs per female within 24 h) are produced at 28 
°C and have a viability of 90 % (De Waele & Wilken, 1990). At 28 °C the life cycle 
from adult to adult can be completed within six to seven days (De Waele & Wilken, 
1990). Greenhouse experiments indicated a 341-fold increase in numbers at 
harvesting (Venter et al., 1991) and a 600-fold increase in vitro on groundnut callus 
tissue after only five weeks (Van der Walt & De Waele, 1989). Numerous generations 
can, therefore, be produced during a single growing season because of the short life 
cycle of D. africanus and favourable soil temperatures in the groundnut-production 
areas, which often exceed 25 °C at a depth of 0 to 30 cm (De Waele & Wilken, 
1990). 
1.2.4 Survival 
In the absence of groundnut D. africanus can survive in low numbers on cotton, 
cowpea, dry bean, grain sorghum, lucerne, lupin, maize, pea, soybean, sunflower, 
tobacco and wheat (Basson et al., 1990). The latter crops are commonly grown in 
South Africa and are often included with groundnut in crop rotation systems (Basson 
et al., 1990). Weeds including cocklebur, feathertop chloris, goose grass, jimson 
weed, khaki weed, purple nutsedge and white goosefoot are commonly found in 
groundnut fields and can also serve as temporary hosts (De Waele et al., 1990 & 
1997). D. africanus can also survive South African winters for at least 28 to 32 weeks 
in hulls left behind in the field after harvesting (Basson et al., 1992). Anhydrobiosis is 
one of the main survival strategies of this nematode (Jones & De Waele, 1990; 
Basson et al., 1993), during which storage time of seed has no negative effect on 
surviving nematodes (Basson et al., 1993). Nematodes that survived in hulls and 
seed can re-infest and damage a subsequent groundnut crop, even from small initial 
population densities (De Waele & Wilken, 1990; Venter et al., 1991; Basson et al., 
1992; Mc Donald etal., 2005). 
1.2.5 Symptoms and histopathology 
Symptoms of D. africanus resemble black pod rot caused by the fungus Chalara 
elegans in irrigated groundnut fields (Labuschagne et al., 1980; Prinsloo, 1980) and 
are similar to those caused by Aphelenchoides arachidis (De Waele et al., 1989), 
5 
which was reported on groundnut seed in Nigeria (Bos, 1977; Bridge et a/., 1977; 
Bridge & Hunt, 1985). 
The initial symptom caused by D. africanus appears at the primary infection site 
located on the peg near the connection point at the base of the pod (De Waele et ai, 
1989; Jones &De Waele, 1990) (Fig. 1.1). 
Figure 1.1. Initial symptoms on groundnut pods caused by 
Ditylenchus africanus infection appears at the 
connection of the peg to the pod. 
The outside tissue infected with D. africanus appear dark brown and corky and brown 
and necrotic on the inside upon removal of the peg (De Waele et ai, 1989). D. 
africanus usually penetrates the hull endocarp through openings at the base of the 
exocarp or at the pod apex (Jones & De Waele, 1990). Infected seed are usually 
shrunken, with dark brown to black micropyles and flaccid testae with darker vascular 
strands (De Waele et ai, 1989). The testae of infected seed can, furthermore, easily 
be removed by gentle rubbing and reveals a distinct yellow discoloration on its inner 
layer (De Waele et ai, 1989). 
Histologically the feeding behaviour of D. africanus causes collapse, malformations 
and cell wall degradation (Venter et ai, 1995). This nematode feeds on the 
parenchyma cells surrounding vascular bundles just below the surface of a pod 
(Jones & De Waele, 1990). At advanced stages of the disease D. africanus-infected 
pods appear dead, with dark brown to black veins (De Waele ef ai., 1989). Feeding 
of the nematodes near or in vascular bundles of the seed testa results in darkened 
6 
veins (Jones & De Waele, 1990). D. africanus does not penetrate the cotyledons but 
do feed on embryos (Jones & De Waele, 1990), causing them to turn olive-green to 
brown (De Waele ef al., 1989) (Fig. 1.2). 
Figure 1.2. Ditylenchus africanus on the cotyledon and embryo 
of an infected groundnut kernel. 
1.2.6 Damage potential and economic importance 
D. africanus is considered to be one of the economically most important plant-
parasites that limit groundnut production locally (Jones & De Waele, 1988; Venter et 
al., 1991; Swanevelder, 1997) since this nematode causes severe losses in 
groundnut crops and income (Mc Donald et a/., 2005). At harvesting 90 % of a D. 
africanus population occurs in the pods, which consist of pegs, kernels and hulls 
(Basson ef al., 1991; Dickson & De Waele, 2005). Penetration of D. africanus at the 
infection site located on the peg near the basis of the pod (De Waele et al., 1989; 
Jones & De Waeie, 1990) causes weakening of the peg and pod connection so that 
pods break off during lifting of the crop (Fig. 1.3) and remains behind in the soil 
(Jones & De Waele, 1990). In heavily infested fields D. africanus might cause losses 
of 40 % to 60 % of pods in this way (Jones & De Waeie, 1988). 
7 
Photo: S. Steenkamp 
Figure 1.3. Groundnut plants are lifted at harvesting. In 
severely-infested Ditylenchus africanus fields, 40 to 
60 % of the pods breaks off and remains behind in 
the soil. 
The main effect of D. africanus on groundnut is qualitative, however (Jones & De 
Waele, 1988 & 1990; De Waele et al., 1989; Mc Donald et al., 2005). Breakdown of 
the hull because of D. africanus damage increases water penetration into the pod 
(Venter et al., 1995) and weakened pods often split open during severe infections 
(De Waele et al., 1997). This breakdown of the hull and split pods result in the 
occurrence of second-generation seedlings (Venter et al., 1995; De Waele et al., 
1997) (Fig. 1.4). 
fctt^fcfc ^ IN* / 
j 
} 
.1 VZ^ 
, Wr ??" ?apv §k I j Z9 i MT 
?W mggpwm w* si TO m 
$W jui •fe \ 
1 
i 
? J 1 
? / m evqa li^fe ?'%v i 
Photo: J. Els 
..^ ^ .j 
Figure 1.4. Groundnut seed and pods infected with Ditylenchus africanus. 
8 
Destruction of the seed testa caused by feeding of D. africanus (Jones & De Waele, 
1990; Venter et a/., 1995), furthermore, leads to leaching of chemical compounds 
that function as inhibitors of seed germination (Svamv & Narasimhareddy, 1977) and 
result in the initiation of growth of the hypocotyls (De Waele et a/., 1997) (Fig. 1.4). 
Feeding of the nematodes near or in vascular bundles of the seed testa also results 
in an unattractive appearance of infected seed (Jones & De Waele, 1990) (Fig. 1.4). 
These symptoms of D. africanus infections have a negative effect on the percentage 
of unsound, blemished and soiled (UBS %) kernels (Venter et a/., 1991; Van der 
Merwe & Joubert, 1992; Mc Donald et a/., 2005) and are highly correlated with the 
number of nematodes found in the testa of the groundnut seed (Venter et a/., 1991; 
Mc Donald et a/., 2005). D. africanus infestations, therefore, can have substantial 
financial implications for a producer (Van der Merwe & Joubert, 1992; Mc Donald et 
a/., 2005). Grading of groundnut consignments in South Africa is specified by law and 
kernels are classified into i) choice edible, ii) standard edible, iii) diverse or iv) 
crushing grade (Anon, 1997). Supply and demand dictate the prices of each grade 
and net gain increases with an increase in kernel grading (Mc Donald et a/., 2005). 
The economic importance of D. africanus is determined by the loss in income from 
infected groundnut consignments, which in turn depends on current prices for each 
grading class (Venter et a/., 1991). 
1.3 NEMATODE MANAGEMENT STRATEGIES ON GROUNDNUT 
Damage threshold levels are reached at the lowest nematode population density that 
is still able to cause a measurable reduction in plant growth or crop yield (Barker & 
Nusbaum, 1971). Control of plant-parasitic nematodes implies the application of a 
single measure to reduce or eliminate nematode pests, which in most cases is 
impossible (Viaene et a/., 2006). On the other hand, nematode management 
combines several different measures in consideration of the whole production system 
to achieve non-injurious or sub-economic threshold levels (Viaene et a/., 2006). 
Nematode management is considered effective when the nematode population 
remains below these damage threshold levels (Ferris, 1978). 
As many current management options are becoming ineffective or unacceptable, 
new acceptable, environmentally-sound strategies must be developed (Barker et a/., 
1994). The rationale behind nematode management is either food or profit and is, 
therefore, driven by resource availability. The ultimate goal of reducing nematode 
9 
numbers and increasing yield quantity and quality at cost-effective levels remain the 
same (Sikora et al., 2005). In the 1990's nematode control measures included the 
use of integrated pest management systems (IPM) that still relied heavily on the use 
of chemical control (Sikora et al., 2005). Globally the majority of land is cultivated by 
smallholder farmers using traditional methods (Altieri, 1984; Van der Merwe et al., 
2001). Because of economic constraints research, therefore, started to focus on low-
input methods (Luc et al., 2005). IPM developed into integrated crop management 
(ICM), concentrating on biological and cultural control methods (Sikora et al., 2005; 
Viaene et al., 2006) or natural pest management (NPM) strategies (Sikora et al., 
2005). 
1.4 CURRENT NEMATODE MANAGEMENT TOOLS IMPLIMENTED FOR LOCAL 
GROUNDNUT PRODUCTION 
1.4.1 Nematicides 
Nematicides have been used since the late 19th century and continue to be an 
important part of nematode management programmes since their primary aim is to 
reduce the number of nematodes invading a crop while increasing yield quantity and 
yield quality (Haydock et al., 2006). Although it is important for a nematicide to 
degrade into harmless compounds and not to persist in the environment, it is 
essential that their efficacy last long enough for efficient nematode control (Haydock 
efa/.,2006). 
Nematicides currently registered locally for use on groundnut include fenamiphos and 
terbufos applied at planting and aldicarb, furfural and oxamyl applied at planting and / 
or at the onset of peg formation (Nel et al., 2007). Nematicides are often used on 
groundnut to prevent damage to the pods later in the season (Sikora et al., 2005) but 
long-term suppression of nematode populations is impossible to achieve with 
chemical control (Starr et al., 2002). Nematicides are often ineffective in sufficiently 
reducing nematode population densities (Haydock et al., 2006), especially those 
such as D. africanus which, because of its high reproduction rate produces more 
than one generation during a single growing season (De Waele & Wilken, 1990). 
Most of these nematicides currently registered on groundnut are, furthermore, 
effective for only eight weeks after application (Nel et al., 2007), while the effective 
control of D. africanus requires a nematicide that remains active for at least 12 weeks 
after application (Basson et al., 1992). 
10 
1.4.2 Cultural and biological management strategies 
Nematicides are often too expensive for most smallholder farmers (Sikora et al., 
2005), therefore the latter rely on traditional methods for groundnut production (Van 
der Merwe et al., 2001). For pest management programmes that exclude chemical 
control, cultural control methods are important alternatives (Viaene et al., 2006). 
Cultural management includes the use of certified seed or nematode-free planting 
material (Viaene et al., 2006). Local production of D. a/r/canus-free, certified seed is 
hampered by factors such as the omnipresence of this nematode in the groundnut 
production areas (De Waele et al., 1989), unpredictable efficacy of nematicides 
under harsh conditions (Mc Donald, 1998) and the unavailability of groundnut 
cultivars resistant to D. africanus (Basson et al., 1991; Van der Merwe & Joubert, 
1992; Venter et al., 1993). Heat and mechanical methods of control are not suitable 
for the treatment of groundnut since its seed is soft, moisture-sensitive and easily 
damaged so that these treatments invariably affect germination (Swanevelder, 1997). 
Biological control is another method that is an important alternative in a pest 
management programme that excludes chemical control (Viaene et al., 2006). 
Although biological control holds some promise (Evans et al., 1993), current 
knowledge on microflora and -fauna is not adequate for the successful 
establishment, promotion or effective suppression of nematode population densities, 
especially over the span of a single growing season (Starr et al., 2002). Reliable and 
effective biological control systems are currently more likely to be limited to 
specialised situations where the environment could be manipulated in order to 
promote biological activity (Sikora et al., 2005) and is, therefore, not adequate in 
keeping nematode populations below damage threshold levels on crops grown in 
most agricultural systems (Viaene et al., 2006). 
1.4.3 Crop-based management strategies 
Crop-based management tools are mainly implemented to achieve high yields and 
improve soil fertility while reducing soil erosion, nematode, insect, disease and weed 
problems (Sikora et al., 2005). Pest control through crop management includes 
starvation and trapping of the pest, antagonism and stimulation of soil antagonistic 
potential and / or biofumigation (Sikora et al., 2005). 
11 
1.4.3.1 Crop rotation 
Crop rotation remains one of the most important tools for nematode management 
(Viaene et a/., 2006). Well-planned rotation systems with other crops can aid in 
production of a high-quality groundnut yield (Swanevelder, 1997). Each production 
system has different requirements, however, and crops used in rotation are planted 
for different reasons (Sikora et a/., 2005). Locally effective management of D. 
africanus with crop rotation is hampered because of this nematode's ability to survive 
in small numbers on many crops other than groundnut, which are often used within 
rotation with groundnut (Basson et a/., 1990). 
1.4.3.2 Resistant cultivars 
The inclusion of resistant cultivars in pest management programmes is often 
preferred over chemical, biological, cultural or regulatory control components (Barker 
et a/., 1994) because host-plant resistance provides the most economical strategy for 
nematode management (Dickson & De Waele, 2005). Resistant cultivars provide 
additional benefits such as sustainability and cost effectiveness. They are 
environmentally benign (Cook & Starr, 2006), while effectively and economically 
managing nematodes on high- as well as low-value cash crops (Dickson & De 
Waele, 2005; Roberts, 2002) and imply little effort or additional cost to the producer 
(Starr et a/., 2002). Low-value crops that cannot support the costs of expensive pest-
management inputs gain most from the planting of resistant cultivars (Fassuliotis, 
1979). In developing countries and low-cash-crop systems, high-yielding, resistant 
cultivars are likely to be the only viable, long-term solution for nematode control 
(Roberts, 2002). To promote the sustainability of resistance, other management 
strategies should be combined with resistant cultivars (Sikora et a/., 2005), especially 
for those that do not express high levels of resistance or tolerance (Roberts, 2002). 
Rotation with resistant plants is the most effective management tool for a number of 
Meloidogyne species parasitising groundnut (Nusbaum & Ferris, 1973; Barker, 1991; 
Rodriguez-Kabana, 1992; Noe, 1998). Groundnut sources expressing resistance to 
M. hapla (Castillo et a/., 1973; Subrahmanyam et a/., 1983), M. javanica (Sakhuja & 
Sethi, 1985), M. arenaria (Simpson & Starr, 2001; Simpson et a/., 2003) and 
Pratylenchus brachyurus (Smith et a/., 1978; Starr, 1984) are currently available. 
Rotation with resistant groundnut cultivars should, however, also be applicable for 
the control of D. africanus (De Waele et a/., 1990). 
12 
1.5 HOST PLANT RESPONSE 
1.5.1 Host sensitivity 
Host sensitivity depends on environmental effects, plant genotype and the number of 
parasites attacking a plant and is measured by the 'tolerant-intolerant' (or sensitive) 
continuum (Cook & Starr, 2006). Tolerant plants experience less yield suppression 
than intolerant plants (Cook & Evans, 1987; Trudgill, 1991; Roberts, 2002). Plants 
expressing extreme tolerance often show no symptoms of infection and are able to 
produce a normal yield (Bos & Parlevliet, 1995). However, tolerant plants usually 
have larger, healthier root systems and tend to allow greater nematode population 
increases (Roberts, 2002). 
In contrast to tolerant plants, sensitive plants often have a smaller root system due to 
nematode injury (McSorley, 1998) and they usually react with relatively severe 
symptom expression e.g. including yield reduction (Bos & Parlevliet, 1995). Smaller 
root systems increase competition among parasites for feeding sites and food 
reserves, which causes a decline in the rate of population increase (Ferris, 1985; 
McSorley, 1998). A hypersensitive plant reacts violently to attacking parasites by 
preventing further spread of infection through prompt death of invaded tissue (Bos & 
Parlevliet, 1995). In some plant-nematode interactions resistance and tolerance are 
under separate genetic control (Evans & Haydock, 1990; Trudgill, 1991) and are 
often inherited independently from each other (Trudgill, 1991). 
1.5.2 Host efficiency 
Host efficiency is determined by the genetic interaction between plant and nematode, 
which is measured by the phenotypic continuum of 'susceptible-resistant' (Cook & 
Starr, 2006). With the exception of high temperatures that may sometimes erode the 
effectiveness of resistance mechanisms, environmental conditions often play a lesser 
role in the expression of host efficiency (Cook & Starr, 2006). Nematode densities 
expressed as number of nematodes per unit available host tissue is the main factor 
affecting host efficiency (Cook & Starr, 2006). A susceptible plant cannot impede the 
growth or development of a parasite (Bos & Parlevliet, 1995), which results in large 
increases in parasite populations, even from low initial densities (McSorley, 1998). A 
resistant host plant on the other hand resists penetration, development, reproduction 
and spread of a parasite (Bos & Parlevliet, 1995). The expression of resistance by a 
13 
host plant, therefore, depends on the plant's ability to interfere with the reproduction 
potential of the parasite (Sikora et al., 2005) and can be described as low, moderate 
or high (reproduction of a parasite is only allowed at trace amounts or does not occur 
at all) (Roberts, 2002). 
A plant is highly resistant when the final population densities (Pf) of a parasite are 
consistently lower than its initial densities (Pi) (Roberts & May, 1986; Windham & 
Williams, 1988) and when the reproduction rate of the parasite is lower than 10 % of 
the reproduction rate on a known susceptible reference (Hussey & Janssen, 2002; 
Timperefa/., 2003). 
1.5.3 Genes expressing resistance 
Resistance to a plant-parasite could be expressed by a single gene (monogenic), a 
few genes (oligogenic) or many genes (polygenic) (Roberts, 2002). Classification of 
genes is based on their phenotypic expression and includes major (large effects) or 
minor genes (small effects) (Roberts, 2002). Simple-inherited, major-gene resistance 
is often preferred because it is easier to identify and to incorporate in back-crossings 
or pedigree programmes using conventional breeding techniques (Roberts, 2002; 
Simmonds, 1991). 
Resistance is classified as vertical (qualitative e.g. race-, pathotype- or biotype-
specific) or horizontal (quantitative e.g. effective against all variants of the pathogen) 
(Van der Plank, 1978). Vertical resistance is controlled by one to three genes while 
horizontal resistance is polygenetically inherited as several minor genes, often with 
an additive effect (Roberts, 2002). The number of genes and their additive effects will 
determine the level of resistance expression (Jones, 1985). Horizontal resistance 
tends to be more durable or is less circumvented as a result of selection pressure on 
a nematode population (Roberts, 2002). 
Expression of resistance is affected by i) genetic constitution of the host plant and 
parasite, ii) environmental effects and iii) virulence status of the nematodes (Roberts, 
2002). Benefits of resistance are best demonstrated in moderately or severely 
infested fields since susceptible cultivars often express a higher yield potential if the 
nematode populations are below damage threshold levels (Sikora et al., 2005). 
Apparent negative effects of resistance on yield are probably due to linkage drag 
whereby genes with negative effects on yield are linked to resistance loci (Cook & 
14 
Starr, 2006). However, no data exist to confirm a direct effect of resistance genes on 
reduced yields (Cook & Starr, 2006). Modern breeding programmes dealing with 
introgression of resistance usually make conscious efforts to increase the yield 
potential of resistant cultivars (Church et al., 2005; Ogallo et al., 1999). 
1.6 MOLECULAR MARKERS 
The regions within genomes containing genes associated with a particular 
quantitative trait are known as quantitative trait loci (QTLs) (Collard et al., 2005). 
Linkage maps are constructed using DNA markers to identify chromosomal regions 
containing genes that control simple and quantitative traits using QTL analysis 
(Collard et al., 2005). DNA markers tightly linked to important genes may serve as 
molecular tools for marker-assisted selection (MAS) in breeding programmes (Ribaut 
& Hoisington, 1998). A locus consists of genetic markers occupying specific genomic 
positions within chromosomes and the development of DNA or molecular markers 
create opportunities to select for QTLs (Collard et al., 2005). 
Use of MAS together with phenotypic selection is more effective, reliable and cost-
effective than conventional plant breeding methodology and is widely accepted as a 
valuable tool for the improvement of crops (Collard et al., 2005). Genetic markers 
associated with M. incognita race 2 (Fourie et al., 2008) and M. arenaria (Tamulonis 
et al., 1997) resistance were identified in soybean for use in MAS and random 
amplified polymorphic DNA (RAPD) analysis resulted in the discovery of three 
markers associated with root-knot nematode resistance in groundnut (Burow et al., 
1996). RAPD is a high-volume technique in which multiple markers can be generated 
from a single DNA preparation (Collard et al., 2005). Church et al. (2001) identified 
the RFLP markers R2430E and S1018E that flanked a dominant-gene locus for root-
knot nematode resistance in groundnut. Although biotechnology is transforming ways 
in which resistance can be incorporated by the use of MAS, it does not eliminate the 
need for verification of the resistant phenotype by direct evaluation of nematode-host 
interaction in the field (Cook & Starr, 2006). 
1.7 RATIONALE AND LAYOUT OF THE CURRENT STUDY 
D. africanus is present throughout the local groundnut production area (De Waele et 
al., 1989) and causes severe losses in groundnut crops and income (Jones & De 
Waele, 1988; Venter et al., 1991; Swanevelder, 1997; Mc Donald et al., 2005). Other 
15 
nematode species have been successfully and economically controlled through the 
use of resistant cultivars (Nusbaum & Ferris, 1973; Barker, 1991; Rodriguez-Kabana, 
1992; Noe, 1998; Roberts, 2002; Starr et al., 2002; Dickson & De Waele, 2005; Cook 
& Starr, 2006). Use of resistant groundnut cultivars may also be applicable for the 
control of D. africanus on groundnut (De Waele et al., 1990). However, no D. 
a/r/'canus-resistant groundnut cultivars are currently available on the market. The 
objectives for this study, therefore, were to: 
i) Identify at least one groundnut genotype with sufficient resistance to D. 
africanus that would also be sustainable under field conditions. 
ii) Compare the reproductive and damage threshold levels of D. africanus on 
susceptible, tolerant and resistant genotypes. 
iii) Establish whether the resistance expressed is present in callus tissue of 
this genotype. 
iv) Establish whether there are differences in the reproduction and damage 
potential of D. africanus from different localities in the groundnut-
production areas of South Africa on resistant genotypes identified in this 
study. 
v) Establish the mechanism of resistance to D. africanus by means of 
histopathlogy. 
vi) Establish the origin of the resistance trait. 
vii) Identify possible molecular markers associated with the resistance trait. 
To achieve the objectives set for this study, the chapters of this thesis consisted of 
the following: 
Chapter 1 provides an overview on the groundnut crop and on D. africanus as a 
plant-parasite on groundnut in South Africa. Aspects discussed include the 
importance of groundnut as a food and income source and the qualitative effects of 
D. africanus on groundnut production. Management tools currently applied or 
available for D. africanus control are also discussed. 
General materials and methods are provided in Chapter 2. Only those specific to 
each chapter were excluded in the latter and provided within the respective chapters. 
Chapters 3 to 6 comprise the investigations done by the author. 
16 
Chapter 3 consists of three parts. In the first part of the study groundnut genotypes 
with D. africanus resistance were identified and the resistance was verified for 
sustainability under field conditions. In the second part of Chapter 3 the reproduction 
rate and damage threshold levels of a range of initial D. africanus population 
densities (Pi) were determined on Sellie, Kwarts, PC254K1 and PC287K5. In the 
third part of Chapter 3 the reproduction rate of D. africanus was studied on callus 
tissue initiated from leaves of Sellie, Kwarts, PC254K1 and PC287K5 to determine 
whether the resistance expressed by PC254K1 will be present in callus tissue. 
Chapter 4 consists of comparative studies done on the reproduction and damage 
potential of five geographically-isolated D. africanus populations representative of the 
groundnut-production areas in South Africa. These studies were done under 
controlled and semi-controlled conditions in growth cabinets, greenhouse and 
microplots to determine differences in reproduction rates and / or temperature 
preferences and to compare the reproduction rates and damage potential of the five 
D. africanus populations on Sellie and PC254K1. 
In Chapter 5 the mechanism of resistance to D. africanus expressed in PC254K1 
was studied to determine histopathological differences associated with PC254K1's 
resistance. 
Chapter 6 comprises a study on the genetics of the resistance identified in Chapter 3 
and a search for molecular markers associated with the resistance trait. The number 
of gene(s) involved in the expression of the resistance trait was determined. 
Molecular markers were mapped and the magnitude of the association between the 
marker and D. africanus resistance was measured. Linkage analysis and drawing of 
the linkage map was done. 
17 
CHAPTER 2 
MATERIALS AND METHODS 
2.1 MICROPLOT FACILITIES 
Microplot trials were conducted in two facilities located on the premises of the 
Agricultural Research Council - Grain Crops Institute (ARC-GCI). Contrary to 
greenhouse conditions, microplots allow for studies of soil-borne pathogens, their 
host plants, abiotic and biotic interactions under semi-controlled environmental 
conditions, i.e. conditions that are more representative of natural environments 
(Abawi & Mai, 1980; Johnson et a/., 1981; Caswell et a/., 1985). Microplots provide 
the additional benefit of the ability to manipulate the substrate as well as certain 
variables such as the introduction of specific nematode species and / or numbers 
(Johnson etal., 1981). 
One microplot facility used in this study (Fig. 2.1) consists of a set of 20 1.1 x 2.1 x 
0.5 m3 rectangular, clay-brick troughs. The troughs are built over a drainage system 
to prevent water logging. Plants are protected against hail by a hail net installed 2 m 
above the plots. 
Figure 2.1. A microplot facility of 20 rectangular brick troughs 
covered with a hail net. 
18 
An irrigation system feeds two evenly-spaced micro-sprayers placed in the centre of 
each plot from a main feeding line, delivering approximately 25 + 4 mm water in 15 
min. 
The other microplot facility (Fig. 2.2) consists of concrete pipes buried vertically in 15 
rows. One row consists of eight, evenly-spaced pipes, each 500 mm deep with a 
diameter of 1 m. 
Figure 2.2. A microplot facility of evenly-spaced, concrete 
pipes and hail-net cover. 
The facility is also built over a drainage system to prevent water logging and a hail 
net, installed + 3m above the plots, protects plants from hail damage. The irrigation 
system feeding this facility consists of micro-sprayer lines connected to a main 
feeding line. Each micro-line feeds a pot through a micro-sprayer placed in the centre 
of the pot and delivers approximately 25 + 4 mm water in 15 min. Irrigation of trials 
conducted in both microplot facilities was supplementary to rainfall. 
2.2 SOIL FOR GREENHOUSE AND MICROPLOT TRIALS 
A sandy-loam soil (Hutton) consisting of 93.6 % sand, 3.9 % clay, 1.9 % silt and 0.6 
% organic material was used in the microplot and glasshouse trials throughout the 
study. The soil was obtained from a farm that is situated 24 km from Leeudoringstad 
(27.26°S, 26.47°E). 
19 
2.2.1 Fumigation of soil 
Soil was fumigated after collection with ethyl dibromide AL (EDB) before filling of the 
microplots or plastic pots used in the greenhouse. EDB used during the current study 
is a registered nematicide and has an active ingredient of 1 800 g per I (Nel ef a/., 
2007). The product was used throughout this study at a rate equivalent to 50 I per 
hectare. Soil fumigation with EDB eliminates undesired organisms that could affect 
the data. EDB was applied manually with a special commercial hand applicator 
(Marunata, Telex 5423339, Mannak J., Kyoto, Japan) that is shown in Figure 2.3. The 
applicator consists of a sealable reservoir, injector handle and dosage control screw 
mounted on one end of a steel shaft. Situated on the opposite end of the shaft is an 
adjustable ring for depth control and release holes at the tip. 
Figure 2.3. A special hand applicator 
used for EDB fumigation of soil. 
Operators wore protective clothing and full-face gas masks during calibration of the 
applicator as well as during application of EDB to the soil. Accurate calibration of the 
applicator is achieved through adjustment of the control screw until the desired 
volume is obtained by each of 10 consecutive injections into a calibrated glass 
measuring cylinder. The required application depth was achieved and maintained by 
adjusting the adjustable ring to the required distance (30 cm) from the tip of the shaft. 
20 
Fumigation of soil for filling microplots or pots was conducted outdoors on a flat-
surfaced area covered with a tarpaulin. Soil destined for fumigation was shovelled 
onto the spread tarpaulin in a layer of at least 30 cm deep to accommodate the part 
of the steel shaft of the applicator that needs to be inserted into the soil. For 
application of EDB the steel shaft was pushed vertically into the soil up to the 
restraining ring (adjusted to an application depth of 30 cm). The injector handle was 
then pushed to release the required dosage rate through the release holes that are 
situated at the tip of the shaft. Applications were done in parallel rows. Injections 
within each row were 30 cm apart and rows were spaced 30 cm apart. Escape of gas 
from the soil was minimised by stepping onto the injection hole with a rubber-soled 
shoe immediately after release of the product into the soil. A second tarpaulin was 
used to cover the freshly fumigated soil to further prevent gas from escaping and to 
prevent contamination of the treated soil. Three weeks after fumigation the soil was 
used to fill microplots or pots for greenhouse trials. 
To minimise the risk of EDB residual effects (Nel ef al., 2007) soil in the microplots 
was re-fumigated (Fig. 2.4) three weeks before planting of each trial. The fumigation 
procedures followed in microplots were similar to those of the soil spread out on a 
tarpaulin described above. 
Figure 2.4. Operators wearing full protective clothing during 
fumigation of soil with EDB in the microplots. 
21 
2.2.2 SOIL NUTRIENTS 
Before each trial was planted a soil nutrient analysis was done by the Soil Science 
Laboratory of the ARC-GCI, Potchefstroom. Nutrients were added based on a soil 
analysis and nutrient guidelines for groundnut production (Swanevelder, 1997). 
Nutrient requirements for each trial will be provided in the respective chapters. 
Nutrients were incorporated into the soil contained in pots (greenhouse) or microplots 
as follows: 
2.2.2.1 Greenhouse 
The soil in each pot was emptied and spread onto a thoroughly cleaned plastic tray. 
The required nutrients were sprinkled over the soil surface and thoroughly mixed with 
the soil by hand. The pot was then refilled with the soil and nutrient mixture. After 
each mixing the trays and gloved hands of the operator were cleaned. 
2.2.2.2 Microplots 
Nutrients were incorporated into the soil in the microplots as demonstrated in Figure 
2.5. First the required amount of nutrients was evenly broadcast over the soil surface 
and then worked into the top 30 cm of the soil with a garden fork. 
Figure 2.5. Nutrients are incorporated into the top 30 cm of 
the soil with a garden fork. 
22 
2.2.2.3 Field sites 
Methods and rates of nutrients applied at the research station at Jan Kempdorp 
(27.95°S, 24.85°E) and on a producer's farm near Hartswater (27.83°S, 24.79°E) 
were based on those used for the respective localities and as recommended after 
soil analyses. The soil type at both localities is Hutton. Herbicides were applied by a 
tractor-mounted implement as and when required (Fig. 2.6). 
Figure 2.6. Application of herbicides with a tractor-mounted 
implement. 
2.3 BIOLOGICAL MATERIALS 
2.3.1 GERMPLASM 
2.3.1.1 Seed 
The germplasm bank of the Groundnut Breeding Unit of the ARC-GCI provided seed 
of groundnut genotypes used in the first microplot trial during the evaluation of D, 
africanus. Seed of genotypes in the second microplot trial that replaced genotypes 
that did not show D. africanus resistance in the first microplot trial (Chapter 3), were 
also obtained from the Groundnut Breeding Unit. However, seed required of the 
breeding lines PC254K1 and PC287K5 and the cultivars Sellie and Kwarts in the 
second microplot trial (Chapter 3) and for the rest of the greenhouse, microplot and 
23 
field trials during this study were produced in the uninoculated section (nematode-
free section) of the first microplot trial (Chapter 3). 
2.3.1.2 Seed treatment 
Pots used in greenhouse trials and the plots in microplot trials were watered a day 
before planting to enhance germination of the seed (Swanevelder, 1997). Standard 
treatments used in groundnut production were adopted for all seed used in the trials 
during this study. The recommended dosage rate of the standard fungicide thiolin is 
based on seed mass (120 g fungicide per 50 kg seed) and it was always applied 
before planting. For thiolin application the relevant seeds were weighed, placed in a 
paper bag and the required rate of the powder-formulation fungicide was added. To 
ensure proper coating of the seeds by the fungicide, each paper bag containing 
seeds and fungicide was shaken rigorously for at least 30 s. Protective gloves and 
face masks were used throughout the process to protect the operator from contact by 
and inhalation of the fungicide. Treated seeds were planted directly afterwards by 
hand to the depth of 5 cm in soil and inter- and intra-row spacing relevant to the 
specific method of groundnut production (Swanevelder, 1997). 
The required amount of Bradyrhizobium arachis nitrogen-fixing bacteria were added 
onto the treated seed before nematode inoculation. The recommended dosage rate 
for the nitrogen-fixing bacteria is 250 g per 50 kg seed. 
2.3.2 NEMATODES 
Microplot, greenhouse and growth cabinet trials were inoculated with nematodes 
extracted from in vitro D. a/r/'canus-groundnut-callus-tissue cultures (Fig. 2.7 A) that 
are maintained in a growth cabinet (Fig. 2.7 B) at a constant temperature of 26 °C 
(Van der Walt & De Waele, 1989). 
24 
Figure 2.7. A. Ditylenchus africanus groundnut callus tissue cultures such as those used for 
the inoculation of microplot and greenhouse trials in this study. 
B. Growth cabinets used for the incubation of Ditylenchus africanus groundnut 
callus tissue cuitures. 
2.3.2.1 Nutrient medium and callus tissue culturing 
The nutrient medium for groundnut callus tissue culturing (Van der Walt & De Waele, 
1989) used during this study was prepared as follows: 4.4 g basal salt mixture, 1 ml 
vitamin B5 complex, 30 g sucrose and 0.9 g caseinhydrolysate were transferred to a 
1-liter glass flask. The flask containing the latter ingredients was topped up with 
distilled water to 1 I. The contents of the flask were mixed thoroughly with a magnetic 
stirrer. During the mixing procedures the pH of the mixture was adjusted to 5.8 using 
t molar HCI / NaOH. Gelrite (8 g) was added to the mixture after having adjusted its 
pH. The mixture was then autoclaved at 120 °C for 15 min. 
From here on were conducted under a laminar flow cabinet in sterile conditions. The 
autoclaved mixture was left to cool down for approximately 10 min before injecting 5 
ml kinetin and 10 ml 2.4-D through millipore filters into the cooled mixture. The latter 
two ingredients completed the medium and were thoroughly mixed with the contents 
by gently shaking of the flask by hand. The medium was immediately poured into 65-
mm petri dishes, each dish receiving approximately 20 ml medium. The medium 
contained in each petri dish was allowed to cool in the laminar flow cabinet for + 20 
min before used for transferring of the callus tissue. 
Callus was propagated on the growth medium described above from young, surface-
sterilised groundnut leaves (Van der Walt & De Waele, 1989) collected from the 
cultivar Sellie planted in a greenhouse for this purpose. Surface sterilisation of the 
25 
leaves proceeded as follows: freshly collected leaves were rinsed under a gentle 
stream of tap water to remove any unwanted materials from their surfaces and then 
transferred to a glass beaker that contained a 70 % ethanol solution. The leaves 
were allowed to soak in the ethanol solution for 30 s. After 30 s the ethanol solution 
was carefully poured off until only the leaves remained in the beaker. A 2-% NaOCI 
solution mixed with two drops of dishwashing liquid was then poured into the beaker 
onto the leaves, which were allowed to soak in the latter solution for 15 min. The 
NaOCI solution was then carefully poured off from the leaves and replaced with 
sterilised, distilled water. The latter rinsing process was repeated four times with 
sterilised distilled water to remove all remaining traces of the NaOCI solution, which 
completed the surface-sterilisation process of the leaves. The sterilised leaves were 
cut on a sterilised glass pane with a sterilised scalpel into 1-cm3 pieces. Each piece 
was transferred from the glass pane with a sterilised pincet onto a growth medium 
contained in a petri dish. The petri dish was then sealed with parafilm. Growth of the 
callus was promoted in darkness for four months in a growth cabinet (Fig. 2.7 B) at a 
constant temperature of approximately 26 °C. 
2.3.2.2 Inoculation of groundnut callus tissue with D. africanus 
All the D. africanus groundnut callus tissue cultures were initiated using nematodes 
extracted from pods (Bolton et a/., 1990) collected from D. africanus-infested fields. 
Extracted nematodes were surface-sterilised (Meyer, 1984) before being inoculated 
on the callus tissue. Each nematode was picked with a sterilised needle from a petri 
dish containing soaked pod tissue and transferred to a 5-ml centrifugal tube that 
contained 2 % streptomycin sulphate. The nematodes were allowed to settle at the 
bottom of the centrifugal tube in the streptomycin sulphate solution for 24 h. After 24 
h the streptomycin sulphate was carefully pippetted off using a fine sterilised pasteur 
pipette and replaced with sterilised, distilled water pipetted onto the nematodes in the 
centrifugal tube. The nematodes were then allowed to re-settle at the bottom of the 
centrifugal tube before repeating this process two more times with distilled water to 
remove all remaining traces of the streptomycin-sulphate solution. 
2.3.2.3 Renewal of D. africanus callus tissue cultures 
Maintenance of D. africanus callus tissue cultures requires renewal of each culture 
every three to four months. Mature D. africanus cultures are presented in Fig. 2.8 A & 
B. Renewal of D. africanus callus tissue cultures was done as described by Van der 
26 
Walt and De Waele (1989). Callus and nutrient medium (Fig. 2.8 B) were cut into 10-
to 20-mm pieces, which contained D. africanus eggs, juveniles and adults. All pieces 
of callus tissue and medium were used to inoculate a fresh callus growing on a 
growth medium (Fig. 2.8 A). Petri dishes containing freshly inoculated callus were 
sealed with parafilm and incubated in darkness at 26 °C \n a growth cabinet, 
Figure 2.8. A. A mature four-month-oid groundnut callus. 
B. A Ditylenchus afr/canus-infested groundnut callus 
tissue culture due for renewal. 
2.3.2.4 Extraction of D. africanus from callus tissue 
Nematodes were recovered from the cultures according to the procedures of De 
Waele and Wiiken (1990). Nematodes present on the lid of the petri dish and on the 
surface of the intact callus tissue and growth medium were rinsed with a slow trickle 
of + 20 ml tap water into a calibrated 500-ml glass beaker. The callus tissue together 
with growth-medium pieces was then sectioned with a scalpel, placed on a 710-^m 
pore sieve nested on a 45-um-mesh sieve and rinsed with tap water. This allowed 
the cells of the callus tissue to separate so that the nematodes within the tissue and 
in the growth medium could be rinsed through the 710-u.m-mesh sieve and collected 
on the 45-um-mesh sieve. These nematodes were washed into the 500-ml glass 
beaker that already contained the nematodes from the petri-dish lid and the surface 
of the callus tissue and growth medium. 
27 
2.3.2.5 Determination of the number of nematodes extracted from callus 
tissue 
The 500-ml beaker containing the extracted nematodes described above was topped 
up with tap water to 300 ml. Nematodes were kept suspended in the water by means 
of a magnetic stirrer to ensure that sub-samples were representative of the nematode 
population in the suspension. Three 10-ml sub-samples of the nematode suspension 
were transferred with a pipette from the beaker into counting dishes while stirring the 
suspension. Eggs, juveniles and adults in each 10-ml sub-sample were counted 
using a research microscope and the number of nematodes present in each counting 
dish was used to calculate the mean number of nematodes per 10 ml nematode 
suspension: (number of nematodes in sub-sample 1 + number of nematodes in sub-
sample 2 + number of nematodes in sub-sample 3) / 3. This figure was used to 
calculate the total number of nematodes present in the 300-ml nematode suspension 
according to the equation y = n x 300 ml / 10 ml where: y = number of nematodes 
per 300 ml and n = mean number of nematodes per 10-ml nematode suspension. 
2.3.2.6 Preparation of nematode inoculum and inoculation procedures 
The desired number of nematodes to be inoculated per plant / plot was always 
suspended in 20-ml aliquots of water. To achieve this the 300-ml nematode 
suspension describe above was diluted to a volume that would yield the desired 
number of nematodes needed to inoculate each plant / plot suspended in 20 ml 
water. When insufficient numbers of nematodes were present in the 300-ml 
suspension another callus culture was put through the extraction process as 
described under 2.3.2.4. Plants were inoculated at planting by pipetting the required 
volume of nematode-suspension evenly over the seeds. 
2.4 DATA COLLECTION AND TRIAL ASSESSMENT PROCEDURES 
2.4.1 Sampling 
2.4.1.1 Greenhouse trials 
The aboveground parts of plants were cut off and discarded. Thereafter every pot 
was emptied separately onto a clean plastic tray so that the whole pod system of 
each plant could be collected from the soil. The pod system was placed in a marked 
28 
plastic bag after removal of the roots from the system. Soil and roots were then 
discarded. Pod samples were kept moisture-free and stored at 4 °C until nematode 
extraction. Nematodes were extracted from the pod tissues within five days of 
sampling. 
2.4.1.2 Microplot and field trials 
For microplot and field trials plants were carefully lifted with a garden fork at 
harvesting, taking care to prevent pods from breaking off in the soil. Aboveground 
parts of randomly selected plants collected for nematode assessments were 
removed and discarded. In the microplot trials whole pod systems of all plants were 
collected and stored as described above. Roots of plants from the field trails were left 
attached to the pod system and together with a 200-ml sub-sample of soil collected 
from the root zone of the plant, placed into a marked plastic bag because nematode 
assessments were also made in root and soil samples to determine nematode 
species other than D. africanus present in the field. Samples were stored at 4 °C until 
extraction, which was within three days of sampling. 
Plants used for yield assessment in microplot and field trials were lifted from the soil 
as described above. The pods were removed and transferred to appropriately 
marked paper bags. The rest of the plants were discarded. The bags were placed in 
a greenhouse for approximately seven days to allow air-drying of the pods at an 
ambient temperature of between 18 and 27 °C. 
2.4.2 Nematode extractions 
2.4.2.1 Soil samples 
Nematodes were extracted from soil samples according to the sugar-flotation method 
described by Jenkins (1964). Soil samples were sub-sampled to 200 cm3 and 
thoroughly mixed with 400 ml tap water contained in a 5-I, calibrated beaker. The 
suspension was poured through a course kitchen sieve into another 5-I container to 
get rid of grit and debris. Then it was stirred and allowed to settle for 30 s. The 
supernatant was then poured through a 750-^m-mesh sieve nested on a 45-^m-
mesh sieve. The residue that remained behind on the 45-p.m-mesh sieve was poured 
into a 50-ml centrifuge tube and was centrifuged. The supernatant was discarded 
29 
and replaced by a 1.13 g / cm3 sucrose solution (624 g sucrose dissolved in 1 I tap 
water), which was thoroughly mixed with the sediment in each tube and centrifuged 
for 1 min at 2 500 revolutions per minute. The supernatant was poured onto a 45-jam-
mesh sieve and residual sucrose rinsed from the nematodes with a gentle stream of 
tap water. Nematodes were then washed into a 130-ml plastic sample flask with + 20 
ml tap water and stored at 4 °C. Nematodes in each sample were counted using a 
research microscope within three days of extraction. Plant-parasitic nematodes other 
than D. africanus collected from the soil samples of the field trials were identified to 
genus level and expressed as nematodes per 200 cm3 soil. 
2.4.2.2 Root and peg samples 
The method used for the extraction of D. africanus from roots and pegs was 
described by De Waele et at. (1987). All root and peg tissues were removed from 
plants sampled from microplot and field trials and cut into 1-cm pieces. These pieces 
were thoroughly mixed by hand and sub-sampled to 5 g. The sub-samples were each 
macerated for 2 min in 250 ml tap water using a high-speed, domestic blender. The 
macerated tissue was poured onto a 750-jj.m-mesh sieve nested on a 45-|am-mesh 
sieve. Course plant material that collected on the 750-jj.m-mesh sieve was discarded 
after it was thoroughly rinsed with tap water and the residue from the 45-|u.m-mesh 
sieve was washed into a 50-ml centrifugal tube. Four centrifugal tubes containing the 
residue from the 45-jj.m-mesh sieve of four 5-g root or peg samples were topped up 
to 50 ml with tap water. Kaolin (2 cm3) was added and thoroughly mixed with the 
water and residue. The four mixtures were centrifuged for 5 min at 2 000 revolutions 
per minute. During centrifugation the kaolin trapped the nematodes at the bottom of 
the tube. The supernatant in each centrifuge tube was discarded. 
Nematodes trapped at the bottom of each tube were separated from the kaolin using 
the sugar-flotation method described previously. Nematodes were washed into a 
130-ml plastic sample flask in + 20 ml tap water. After storage at 4 °C in a fridge the 
water containing the nematodes was poured into a counting dish and specimens 
were counted using a research microscope within three days of extraction. Eggs 
were not included since the developing juveniles in the eggs could not be identified. 
Nematodes were expressed as number per 5 g roots or 5 g pegs. 
30 
2.4.2.3 Hull and kernel samples 
Pods of plants collected for nematode assessments were removed by hand from 
lifted plants. The pods were mixed and 20 were randomly picked. Hull and kernel 
tissue was extracted separately according to the procedures described by Bolton et 
al. (1990). Hulls were broken by hand and kernels were cut with a scalpel into small 
pieces to facilitate migration of the nematodes from the plant tissue into the water. 
The tissue was soaked for 24 h in petri dishes containing + 20 ml tap water. After 24 
h the contents of each petri dish were poured onto a 750-|j.m-mesh sieve nested on a 
45-|u.m-mesh sieve. Course plant material collected on the 750-|j.m-mesh sieve was 
discarded after thoroughly washing it through with tap water. The nematodes 
collected during this process on the 45-|u.m-mesh sieve were washed into a 130-ml 
plastic sample bottle with + 20 ml water. After storage at 4 °C the contents of each 
sample bottle were poured into a counting dish and the nematodes counted using a 
research microscope within three days after extraction. Nematode numbers were 
expressed per 5 g hulls or 5 g kernels. Numbers of D. africanus present per 15 g 
pods (Pf) was calculated by adding the number of D. africanus per 5 g pegs, 5 g 
kernels and 5 g hulls. 
2.4.3 Crop yield 
2.4.3.1 Yield quantity 
To determine yield mass per replicate the procedures described by Mc Donald 
(1998) were followed. Pods collected from plants sampled for yield assessments 
were weighed and the pod mass for each replicate was recorded. A sub-sample of 
500 g pods per replicate was shelled and the kernels were used to determine yield 
quality. 
2.4.3.2 Yield quality 
The quality of each kernel-yield replicate was determined in a 200-g kernel sub-
sample obtained from the shelled 500-g pod samples described above. These sub-
samples were subjected to grading procedures stipulated by the Act on Agricultural 
Product Standards, 119 of 1990 (SA, 1997). Each 200-g sub-sample was placed on 
sieves nested on each other in receding order of aperture size. A solid pan was 
31 
placed underneath the 7.20-mm-aperture slotted sieve. Kernels were sifted through 
6.75-mm, 6.00-mm and 7.20-mm-aperture slotted sieves, similar to those used in the 
groundnut industry. The kernels remaining on the 6.75-mm-aperture and 6.00-mm-
aperture sieves were sorted into unsound, blemished, soiled, damaged, shrivelled 
and sun-dried categories. Kernels on the 7.20-mm-aperture sieve were sorted into 
unsound, blemished and unsplit. Standards used for the grading of kernels are 
specified as follows: 
Blemished kernels: Whole kernels with coloured blotches or streaks that 
are not associated with the specific groundnut genotype. 
Damaged kernels: Broken or whole kernels of which the testa are loose, 
cracked or have partially or entirely come off. 
Mould-infested kernels: Those kernels on which external or internal mould 
growth is visible with the naked eye or those kernels damaged or 
discoloured by mould growth. 
Shrivelled kernels: Whole kernels with a shrivelled, grooved or dented 
appearance. 
Soiled kernels: Whole kernels that are soiled to such an extend that their 
appearance is affected. 
Sun-dried kernels: Those kernels of which the testa comes off easily 
when lightly rubbed by hand. 
Unsound kernels: Mould-infested kernels or kernels that are decayed, 
chalky, have sprouted or are discoloured and are not characteristic of 
healthy kernels of the specific genotype. 
The different grades of kernels from each sieve were weighed separately and 
evaluated as stipulated by the abovementioned products standard act (Table 2.1). 
Table 2.1. Local specifications for groundnut grading as stipulated by the Act on Agricultural 
Products Standards, 119 of 1990 (SA, 1997). 
Grade Sieve slot Seed on sieve UBS Unsound Total defects 
(mm) (%) (%) (%) (%) 
Choice 6.75 >20 <10 15 <20 
Standard 6.75 >10 <20 <10 <35 
Diverse 6.00 >40 <30 <15 <35 
Crushing 7.20 >20 >30 >15 >35 
32 
2.5 STATISTICAL ANALYSES 
Degrees of freedom (D.F.) (error) > 18 (Van Ark, 1981) were pursued in all trials. 
Data were entered on a spreadsheet of Microsoft Excell before being imported into 
Stat Graphics 5 Plus for Windows. The appropriate analysis of the nematode data 
will be discussed in each relevant chapter. Nematode data were ln(x+1) transformed 
before statistical analysis unless stated otherwise. Final nematode number (Pf) per 
15 g pods was used as indicators of resistance or susceptibility of each groundnut 
genotype because D. africanus populations peak at harvest time in pods (Basson et 
al., 1990). Unless otherwise stated nematode numbers from root and soil samples 
were not determined because D. africanus reproduces poorly in roots (Van der Walt 
& De Waele, 1989; Basson et al., 1990 & 1991; De Waele et al., 1990; Jones & De 
Waele, 1990) and numbers in the soil are small and variable (Venter et al., 1992). 
The reproduction factor (RF) of D. africanus on each groundnut genotype was 
determined in all the trials using Oosterbrink's equation: RF = final population density 
(Pf) / initial population density (Pi) (Windham & Williams, 1987). The RF of D. 
africanus on groundnut genotypes was determined in most trials because it provides 
a basic measurement for the reproduction of a nematode population on a crop 
(Windham & Williams, 1987; Marin et al., 1999) and supports Pf data. The Pi and Pf 
used in the equations were not ln(x+1) transformed because measurements (Pi and 
Pf) of a variable (RF) must be in a nominal scale (Van Ark, 1981). 
Yield mass, yield quality and income for microplot trials as specified in the relevant 
chapters were calculated using Maksi Plan, which is a computer programme 
developed by the ARC-GCI for the evaluation of crop cultivars and provides functions 
and evaluations specifically applicable to this study. Calculations of Maksi Plan were 
based on yield quantity and quality data determined for each genotype and the 
grades were priced according to the local market value for groundnut at the time of 
the study. 
33 
CHAPTER 3 
IDENTIFICATION OF GROUNDNUT GENOTYPES WITH RESISTANCE TO 
DITYLENCHUS AFRICANUS 
3.1 INTRODUCTION 
Currently available control measures are not adequate under all conditions to keep 
D. africanus numbers below damage-threshold levels (Basson et a/., 1990 & 1993; 
Mc Donald et a/., 2005). This nematode is difficult to control because of its ability to 
survive in the absence of groundnut (Basson et a/., 1990; De Waele et a/., 1990 & 
1991; Swart & Jones, 1994), its high reproductive potential and its short life cycle (De 
Waele & Wilken, 1990). These characteristics enable the nematode to quickly build 
up to large populations that cause severe damage (De Waele & Wilken, 1990; De 
Waele et a/., 1990; Venter et a/., 1991; Basson et a/., 1990; 1992 & 1993, Mc Donald 
efa/., 2005). 
Cultivation of resistant crops or cultivars is one of the easiest ways to achieve a 
reduction in initial nematode densities (Brown, 1987) and it provides an effective 
alternative for the management of various plant-parasitic nematodes. Resistant plant 
material is able to resist ingress, establishment and spread of the parasite itself (Bos 
& Parlevliet, 1995) and keep the latter below damage-threshold levels (Roberts & 
May, 1986; Fourie et a/., 1999; Ogallo et a/., 1999; Timper et a/., 2003; Todd et a/., 
2003; Fourie, 2005). The same principle should be applicable to D. africanus 
management on groundnut (De Waele etal., 1990). 
For the successful implementation of IPM programmes, however, it is also important 
to obtain information on the minimum nematode population densities that cause 
measurable damage (damage-threshold levels) on a crop (Barker & Nusbaum, 
1971). If a nematode population could be kept below this damage-threshold level, the 
particular control measure is considered successful (Kim & Ferris, 2002). Damage-
threshold levels of several plant-parasitic nematode species have been studied on a 
wide variety of crops (Ferris, 1978 & 1985; Niblack et a/., 1986b; Koenning, 2000; 
Kim & Ferris, 2002; Perez et a/., 2003; Fourie, 2005; Mc Donald et a/., 2005). 
Greenhouse studies showed that damage by D. africanus on groundnut could 
become significant at an initial population density (Pi) of 50 nematodes (Venter et a/., 
34 
1991), while microplot studies indicated that a Pi of 250 to 750 nematodes will lead to 
downgrading of the cultivar Sellie to standard grade (Mc Donald et ai, 2005). 
Currently there are no cultivars available that have resistance to D. africanus. 
Previously more than 600 genotypes were evaluated for resistance to this nematode 
but none proved useful (Basson et ai, 1991; Van der Merwe & Joubert, 1992). 
Consequently, no information is available in the literature on the comparative effect of 
increments of Pi of D. africanus on susceptible, tolerant or resistant groundnut 
genotypes. The ultimate challenge for any breeding-programme is, furthermore, to 
release new cultivars through the shortest route possible. One possible approach 
could be to reduce the period of microplot and field evaluations by using callus tissue 
to screen genotypes against D. africanus. The objectives of this part of the study 
were to i) identify groundnut genotypes with resistance to D. africanus that would 
also be sustainable under field conditions, ii) compare the reproduction and damage-
threshold levels of D. africanus on resistant, susceptible and tolerant groundnut 
genotypes in microplots and iii) determine whether resistance expressed in pods of a 
resistant genotype is transferred to its callus tissue under controlled conditions. 
3.2 MATERIALS AND METHODS 
3.2.1 Identification of D. afr/camvs-resistant groundnut genotypes 
To achieve the first objective selected groundnut genotypes were evaluated over two 
consecutive seasons in microplot and field trials. The genotypes selected for 
evaluation for D. africanus resistance are listed in Table 3.1. The genotypes were 
selected on the basis of characteristics such as high oleic-acid contents or disease 
resistance (Cilliers et ai., 2001). Kernels of genotypes with a high oleic-acid content 
are less reactive with oxygen and consequently have a longer shelf life (Isleib et ai., 
1994), while the use of genotypes that have resistance to one or more diseases can, 
potentially, reduce input costs for producers (Dickson & De Waele, 2005). 
35 
Table 3.1. Origin and preferred characteristics of groundnut genotypes that were selected for 
the identification of resistance to Ditylenchus africanus. 
Genotype Origin Source Preferred characteristic 
*Sellie Locally bred ARC-GCI1 D. afr/'canus-susceptible standard 
*Kwarts Locally bred ARC-GCI D. afr/canus-tolerant standard 
*Harts Locally bred ARC-GCI resistant to black pod rot 
*UF85 USA Unknown high oleic acid content 
*JL24 Congo SARCCUS2 resistant to tomato spotted wilt virus 
*CG7 Malwi ICRISAT3 resistant to a variety of diseases 
"453 Senegal Unknown high oleic-acid content 
"73-30 Senegal Unknown high oleic-acid content 
"PC223 Locally bred ARC-GCI high oleic-acid content 
"PC299K5 Locally bred ARC-GCI high oleic-acid content 
"PC254K1 Locally bred ARC-GCI high oleic-acid content 
"PC287K5 Locally bred ARC-GCI suspected resistance to D. africanus 
*Cultivar 
"Breeding line 
Agricultural Research Council-Grain Crops Institute, Potchefstroom, North 
West Province 
2Southem African Regional Commission for the Conservation and Utilisation of Soil 
international Crops Research Institute for the Semi-Arid Tropics 
3.2.1.1 Trial layout 
3.2.1.1.1 Seed and genotypes 
Seed of the genotypes that were tested in this study were obtained from the 
germplasm bank of the groundnut-breeding department or from nematode-free 
nurseries (Chapter 2). Before planting 5-g sub-samples of the seed of each genotype 
were soaked in water for 24 h (Bolton et al., 1990) to ascertain that the seed was D. 
a/ncani/s-free. The rest of the seed was treated with fungicide and inoculated with 
Bradyrhizobium arachis nitrogen-fixing bacteria (Chapter 2) before planting. 
The groundnut genotypes tested in each microplot or field trials during each season 
are listed in Table 3.2. The cultivar Sellie served as D. a/ncani/s-susceptible standard 
(Mc Donald, 1998) in all trials. Kwarts, classified as tolerant to D. africanus (Mc 
36 
Donald, 1998) was used as the other standard because no source of resistance to D. 
africanus was available at the onset of this study. 
Table 3.2. Groundnut entries for the identification of resistance to Ditylenchus africanus in 
the respective microplot and field trials conducted over two consecutive growing 
seasons (2003-2004 - 2004-2005). 
Microplot trial 1 
(2003-2004) 
Microplot trial 2 
(2004-2005) 
Field trials (2) 
(2004-2005) 
Sellie ' Sellie Sellie 
Kwarts 2 Kwarts Kwarts 
UF85 JL24 JL24 
73-30 Harts Harts 
453 CG7 CG7 
PC223 PC299K5 PC299K5 
PC254K1 PC254K1 PC254K1 
PC287K5 PC287K5 PC287K5 
D. a/r/'ca/ws-susceptible standard 
2 D. a/r/canivs-tolerant standard 
3.2.1.1.2 Microplot trials 
The two microplot trials were planted in a facility that consists of a row of 20 
rectangular clay-brick troughs (Chapter 2). The 20 troughs were each filled with 
fumigated, sandy-loam soil (Chapter 2). The soil was fumigated once more in the 
microplots three weeks before planting with EDB as described in Chapter 2. Nutrients 
were added to each trough according to the procedures described in Chapter 2. 
Requirements for the 2003-2004 microplots were 150 g dolomitic lime, 130 g super 
phosphate (10.5 % P), 40 g sodium chloride (KCI) and 35 g calcium nitrite (CaN03) 
per trough. Additional nutrients required for the microplot trial during 2004-2005 were 
200 g dolomitic lime, 139 g super phosphate (10.5 % P), 42 g KCI and 40 g CaN03 
per trough. 
The trials were planted in a randomised complete split-plot design. Factor 1 
consisted of 10 plots artificially inoculated with D. africanus and 10 uninoculated 
plots. The inoculated section received + 3 000 D. africanus of various life stages per 
plant at planting. The nematodes used for inoculation were extracted from in vitro D. 
37 
africanus cultures prepared for inoculation according to procedures described in 
Chapter 2. Factor 2 consisted of the relevant eight genotypes evaluated during each 
season (Table 3.2). Each genotype, inoculated and uninoculated was replicated five 
times. 
The trial layouts were the same in 2003-2004 and 2004-2005, although the 
randomisation in each block was different in the two seasons. Four rows were 
planted in each trough with an inter-row spacing of 45 cm (Chapter 2). Each plant 
row contained 20 seeds of each respective genotype. Seed was planted to a depth of 
5 cm using an intra-row spacing of 5 cm (Chapter 2). 
3.2.1.1.3 Field trials 
Sustainability of resistance of these genotypes under field conditions was tested in 
two separate field trials. Fields naturally infested with plant-parasitic nematodes were 
selected and were used as sites for the trials. One field trial was planted on an 
overhead-irrigated site near Hartswater (27.83°S, 24.79°E) and the other trial on a 
flood-irrigated site located on the ARC Research Station (Fig 3.1) near Jan 
Kempdorp (27.95°S, 24.85°E). 
Figure 3.1. A groundnut field (rial at Jan Kempdorp. 
No D. africanus was artificially added to the soil since both sites were previously 
planted with groundnut and were naturally infested with this nematode. Soil at these 
sites was not treated with any nematicide or fumigant at or after planting of the trials 
38 
because of the risk of contamination of adjacent plots by chemical treatments, 
particularly under irrigation, which could interfere with results (Hough & Thomason, 
1975). Nematode-free controls were, therefore, not included in the field trials. 
The trials were planted in a randomised complete block design (RCBD) including 
eight treatments (genotypes listed in Table 3.2) and six replicates per treatment. 
Each replicate consisted of eight 1-m rows planted at an inter-row spacing of 45 cm 
(Chapter 2). Each row was planted to 20 seeds of each respective genotype. The 
seeds were planted to a depth of 5 cm with intra-row spacing of 5 cm (Chapter 2). 
3.2.1.2 Collection of nematode and yield data 
Nematode and yield assessments were done at harvesting. Four randomly selected 
plants from each row were collected for nematode extraction from the inoculated 
section of the microplot and from all plots in the field trails. Extractions were also 
made from four randomly selected plants from the uninoculated section of the 
microplot trials to confirm that this section remained nematode-free. 
Separate extractions were done from peg, hull and kernel samples from the microplot 
trials and from soil, root, peg, hull and kernel samples from the field trials, following 
the procedures described in Chapter 2. Nematodes were counted using a research 
microscope and the numbers (Pf) were added and expressed as number of D. 
africanus per 15 g pods (Chapter 2). 
In the field trials root and soil samples were included because assessments were 
made of all the nematode species that occurred along with D. africanus. Nematodes 
were extracted from soil and plant tissues according to the methods described in 
Chapter 2. Two different methods were used for extraction of nematodes from root 
samples collected from the field trials. The maceration method (De Waele ef a/., 
1987) was used to extract plant-parasitic nematodes other than Meloidogyne spp. 
from root samples, which were expressed as nematodes per 5 g roots (Chapter 2). 
However, the latter method is not effective for extracting eggs and second-stage 
juveniles (J2) of Meloidogyne spp. from root samples. An additional NaOCI method 
(Riekert, 1995) was used for the extraction of root-knot nematode eggs and J2 from 
root samples, which were expressed as Meloidogyne spp. per 50 g roots. The NaOCI 
method provided a more reliable estimate of the Meloidogyne spp. population 
39 
densities present in the field. All plant-parasitic nematodes other than D. africanus 
were identified to genus level. 
The 16 remaining adjacent plants in each row that were not used for nematode 
extractions were collected for yield assessment in the microplot as well as field trials. 
Yield quality (microplot and field trials) and quantity (microplot trials) assessments 
were made as described in Chapter 2. Yield of the genotypes in the uninoculated 
section of the microplot trials served as nematode-free control to distinguish between 
resistant and tolerant plant responses. 
3.2.2 Reproduction and damage-threshold levels of D. africanus on resistant, 
susceptible and tolerant groundnut genotypes 
The microplot facility used for the second part of the study during 2004-2005 consists 
of concrete pipes arranged vertically in 15 rows of eight pipes each (Chapter 2). The 
120 plots were each filled with the same fumigated, sandy-loam soil (Chapter 2) that 
was used to fill the microplots in the evaluation of the groundnut genotypes against 
D. africanus. Procedures for the fumigation of the soil with EDB are described in 
Chapter 2. The nutrients incorporated into the soil of each plot (Chapter 2) at the 
beginning of the 2004-2005 growing season were 78 g dolomitic lime, 47 g 
superphosphate (10.5 % P), 31 g KCI and 20 g CaN03. 
To accommodate the total number of treatments (192) in the 120 available plots, a 
0.4 x 1 m2 piece of rigid, 1.5-cm thick rubber mat was installed vertically along the 
diameter of each plot after application of nutrients to the soil. The rubber mats 
divided each pot in equal halves and served as a physical barrier for water and 
nematodes between the two halves. Each plot halve therefore served as a separate 
plot. Each plot was planted with six seeds of a groundnut genotype. The seed was 
planted to a depth of 5 cm (Chapter 2). 
Seeds of Sellie, Kwarts, PC254K1 and PC287K5 were obtained from nematode-free 
seed nurseries (Chapter 2). Before planting 5-g sub-samples of the seed of each of 
the groundnut genotypes were soaked in water for 24 h (Bolton et a/., 1990) to 
ascertain that the seed was free of D. africanus infection. The rest of the seed was 
then treated with fungicide and B. arachis as described in Chapter 2. 
40 
The trial was laid out in a RCBD. The four groundnut genotypes were each 
inoculated (Chapter 2) with 0, 50, 250, 1 000, 2 500, 5 000, 10 000 or 20 000 
nematodes (Pi) that originated from Hartswater per plant at planting. Each Pi for each 
genotype was replicated six times. A plastic marker placed in each plot indicated the 
respective genotype, the Pi and replicate number. The trial received supplementary 
irrigation as described in Chapter 2. 
Nematode and yield assessments were made at harvesting. Two plants per plot were 
randomly picked for extraction of nematodes from peg, hull and kernel samples 
(Chapter 2). The nematodes were counted using a research microscope and 
expressed as D. africanus per 15 g pods (Pf) (Chapter 2). The four plants remaining 
per replicate were used for yield quantity and quality assessments, following the 
procedures described in Chapter 2. 
3.2.3 Expression of D. africanus resistance in callus tissue 
To achieve the third objective of this study the reproduction of D. africanus was 
studied on callus tissue from Sellie, Kwarts, PC254K1 and PC287K5 in a growth 
cabinet trial during September and October 2008. The trial was laid out in a RCBD 
with the four different genotypes as treatments, each replicated six times. The callus 
tissue was initiated from young groundnut leaves from each genotype as described in 
Chapter 2. Each six-week old callus was inoculated with 10 adult D. africanus, 
consisting of five males and five females extracted from an in vitro D. africanus 
culture that originated from Hartswater (Chapter 2). Each petri-dish containing the 
inoculated callus was sealed with parafilm. Dishes were marked appropriately to 
indicate the relevant genotype and replicate number. The cultures were incubated in 
darkness for four weeks at 28 °C in a growth cabinet to allow the nematodes to 
complete a minimum of four life cycles (De Waele & Wilken, 1990). The growth 
cabinet was set at a constant temperature of 28 °C and was allowed to stabilise for 
three days before the onset of this trial. After four weeks the nematodes were 
extracted from each callus according to the method described in Chapter 2. The 
nematodes were counted using a research microscope and the numbers were 
expressed as D. africanus per 1 g callus tissue. 
41 
3.3 STATISTICAL ANALYSES 
Unless otherwise stated nematode data were ln(x+1) transformed before being 
subjected to statistical analysis. Nematode data from the microplot and field trails on 
D. africanus resistance and that of the growth cabinet trial were subjected to an 
analysis of variance (ANOVA) (Stat Graphics 5 Plus for Windows). Means were 
separated by an LSD test (P £ 0.05). Nematode data from the microplot trial on the 
reproduction and damage threshold of D. africanus were subjected to linear and non 
linear regression analyses (Genstat for Windows). 
The following nematode variables were determined: Final population densities (Pf) of 
D. africanus in pods and callus tissue of all trials. The RF of D. africanus on each 
groundnut genotype was determined for the microplot and growth cabinet trials as 
described in Chapter 2. RF values were not determined for field trials since Pi, which 
is a factor used in the determination of the reproduction factor, cannot be accurately 
estimated under field conditions (De Waele et a/., 1990; Venter et a/., 1991; Basson 
et ai, 1992; Taylor et a/., 2000). 
The percentage reproduction rate (RR) of D. africanus populations was determined 
on pods from each groundnut genotype from the microplot and field trials on D. 
africanus resistance. Although resistance is a relative concept, RR can be used to 
indicate the level of resistance expressed by the host (Timper et ai, 2003) in the 
absence of an RF value in the field because Pi is not required as a factor to 
determine RR. The RR of each treatment from the inoculated microplot section as 
well as the field trials was calculated using the equation RR = Pf (genotype) / Pf 
(Sellie) x 100. Pi and Pf used in the equation to determine RR were not ln(x+1)-
transformed since measurements (Pi and Pf) of the variable (RR) must be in a 
nominal scale (Van Ark, 1981). 
Yield quantity of cultivars and breeding lines were determined to be used in Maksi 
Plan (Chapter 2) but were not compared with one another in either the microplot or 
field trials on D. africanus resistance or in the microplot trial on reproduction and 
damage threshold levels because yield is determined by intrinsic agronomic traits 
(Fourie, 2005). Quantity data of genotypes evaluated for resistance to D. africanus in 
the field were not shown because yield data of the genotypes was not used in any 
additional calculation to determine the effect of D. africanus on their yields. Quantity 
of the yields produced by the four genotypes at each Pi level in the microplot trial on 
42 
reproduction and damage threshold levels was compared within each genotype to 
determine the effect of this nematode on yield quantity. 
Yield quality of each genotype collected from all of the microplot and field trials was 
determined according to standard grading procedures (Chapter 2). Yield quantity and 
income produced by each genotype in the inoculated as well as the uninoculated 
sections of the microplot trials on D. africanus resistance and for those at different Pi 
levels in the microplot trial on reproduction and damage threshold levels were 
determined by means of Maxi Plan (Chapter 2). For the microplot trials on D. 
africanus resistance yield quantity per hectare for each genotype was calculated 
according to the equation Py = (Yui-Yi) 100 / Yi, where Py = yield response, Yui = 
yield in the uninoculated section and Yi = yield in the inoculated section (Weaver et 
al., 1985). Income difference was calculated using the same equation but replacing 
yield with income: Pin = (lui-li) 100 / li, where Pin = potential income, lui = income 
calculated for the uninoculated section and li = income calculated for the inoculated 
section. Loss or gain of yield and income per hectare of each genotype was then 
compared among the genotypes. 
3.4 RESULTS 
3.4.1 identification of D. afr/'canus-resistant groundnut genotypes 
3.4.1.1 Microplot trials 
3.4.1.1.1 Final nematode population densities, reproduction factor and 
reproduction rate 
Genotypes that showed resistance to D. africanus during the first microplot trial 
during 2003-2004 were re-evaluated in the second microplot trial during 2004-2005. 
Genotypes that did not show any resistance to D. africanus in the first microplot trial 
were replaced by different ones. During both seasons the nematode numbers in the 
pods of Sellie, Kwarts, PC254K1 and PC287K5 varied in spite of a constant inoculum 
rate of 3 000 nematodes per plant in the microplot trials. Significantly lower Pfs were 
extracted from pods of PC254K1 and PC287K5 compared to those of the rest of the 
genotypes during 2003-2004 (Fig. 3.2). 
43 
10 - LSD (P< 0.05) 
8 
! 
o> 6 [
 ll l I I I 
i < 
2 
0 
: I I 
s <o lO v 
O o 0- Q. 
Genotype 
Figure 3.2. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from the inoculated section of a microplot trial during 
2003-2004 (P < 0.05; F-ratio = 10.02). 
12 LSD (P< 0.05) 
10 I I I I I 
CO 
E 8 
CL I I 
S 6 
+ I 
a: 4 -
2 -
0 _ 
TO 
i2 ,-H it: 
o l5 * 
< S to V CM CM o O o 
CL Q. 0. 
Genotype 
Figure 3.3. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from the inoculated section of a microplot trial during 
2004-2005 (P < 0.05; F-ratio = 40.69). 
During 2004-2005 the latter two genotypes along with CG7 maintained significantly 
lower Pfs compared to those of the rest of the genotypes (Fig. 3.3). The Pf of CG7 
was significantly lower than those for the other genotypes tested during 2004-2005. 
44 
PC299K5 also had a significantly lower Pf than those of the susceptible checks Sellie 
and Harts during the same season. 
The relatively low Pf present in pods of genotypes in the inoculated section of the 
microplot trial during 2003-2004 also translated in their respective RF, which were all 
lower than one (Table 3.3). In spite of the low levels during this season the Pf of 
PC254K1 was 53.9-fold lower than that of Sellie. During the same growing season 
the Pf of PC287K5 was 7.2-fold lower than that of Sellie. The RR of PC254K1 and 
PC287K5 relative to Sellie was also very low. Those of the tolerant Kwarts were less 
than a quarter of the RR of Sellie, while the RR of the lines 453 and PC233 were less 
than half that of Sellie. 
Table 3.3. Ditylenchus africanus numbers (Pf), reproduction factors (RF) and reproduction 
rates (RR) in pods of eight groundnut genotypes from a microplot trial during 
2003-2004. 
Genotype Pf(15gpods) RF1 RR" 
UF85 2 700 0.90 100.3 
Sellie3 2 693 0.89 100 
73-30 1 742 0.58 64.7 
PC233 1 335 0.45 49.6 
453 848 0.28 31.5 
Kwarts4 610 0.20 22.7 
PC287K5 372 0.12 13.8 
PC254K1 50 0.02 1.9 
1RF = Pf/Pi 
2RR = Pf (genotype) / Pf (Sellie) x 100 
3D. a/ncanus-susceptible standard 
4D. africanus-tolerant standard 
During 2004-2005 the Pf's overall were much higher than the previous season in the 
microplots (Table 3.4). The Pf in pods of CG7 was 103-fold lower, 26.9-fold lower in 
PC254K1 and 13.4-fold lower in PC287K5 than that in Sellie. The RF's of D. 
africanus were higher than one for the majority of the genotypes. CG7 and PC254K1, 
PC287K5 had RF < 1. The RR's of D. africanus (Table 3.4) for CG7, PC254K1 and 
PC287K5 were 1 %, 3.7 % and 7.5 %, respectively of that for Sellie. The RR of 
tolerant Kwarts was only more than half of that of Sellie, despite the much higher Pf 
in this trial. 
45 
Table 3.4. Ditylenchus africanus numbers (Pf), reproduction factors (RF) and reproduction 
rates (RR) in pods of eight groundnut genotypes from a microplot trial during 
2004-2005. 
Genotype Pf(15gpods) RF1 RR2 
SellieJ 28 116 9.37 100 
Harts 25 649 8.55 91.2 
JL24 15 191 5.06 54.0 
Kwarts4 14919 4.97 53.1 
PC299K5 7 843 2.61 27.9 
PC287K5 2 095 0.70 7.5 
PC254K1 1 046 0.35 3.7 
CG7 273 0.09 1.0 
1RF = Pf/Pi 
2RR = Pf (genotype) / Pf (Sellie) x 100 
3D. afr/canus-susceptible standard 
4D. africanus-to\erani standard 
3.4.1.1.2 Yield assessments 
The difference in yield quality in the inoculated and corresponding uninoculated 
sections of the microplot trials varied between genotypes and over seasons (Tables 
3.5 & 3.6). The UBS % of most genotypes in the inoculated section was generally 
higher than those in the corresponding uninoculated section during both seasons. 
During 2003-2004 PC254K1, PC287K5 and Kwarts (tolerant) had UBS % lower than 
10 and were comparable between the inoculated and uninoculated sections in this 
regard (Table 3.5). Greater reduction in UBS % from nematode inoculated to 
uninoculated were only apparent in PC223, Sellie, UF85 and 73-70. The grades from 
choice to crushing corresponded with the trend in UBS % (Table 3.5). Choice grade 
kernels were obtained from the inoculated and the uninoculated sections of 
PC254K1, PC287K5 and Kwarts. The lines 453 and 73-70 produced standard grade 
kernels in the inoculated and uninoculated sectiond, while PC233, Sellie and UF85 
only yielded standard grade kernels for the uninoculated section. Only diverse and 
crushing grade were obtained from the inoculated sections of the rest of the 
genotypes. 
46 
Table 3.5. Yield quality of eight groundnut genotypes in a microplot trial during 2003-2004. 
UBS Choice Standard Diverse Crushing 
Genotype (%) (%) (%) (%) (%) 
r Ul2 I Ul I Ul I Ul I Ul 
PC223 25 12 0 0 0 66 73 18 27 16 
Sellie3 23 14 0 0 0 68 79 21 21 11 
UF85 22 2 0 0 0 14 76 21 24 65 
73-70 21 12 0 0 46 61 22 21 32 18 
453 15 14 0 0 47 56 18 23 35 21 
Kwarts4 6 5 75 80 0 0 12 9 13 11 
PC287K5 1 4 48 31 0 0 36 33 16 36 
PC254K1 0 0 64 64 0 0 15 9 21 27 
inoculated 
2Uninoculated 
3 D. afr/ca/ws-susceptible standard 
4D. afr/ca/ws-tolerant standard 
Table 3.6. Yield quality of eight groundnut genotypes in a microplot trial during 2004-2005. 
UBS Choice Standard Diverse Crushing 
Genotype I :%) < :%) I :%) (%) (%) 
r Ul2 I Ul I Ul I Ul I LI I 
Harts 26 20 0 0 0 0 66 68 34 32 
PC299K5 21 20 0 0 0 0 52 49 48 51 
Sellie3 20 9 0 0 0 33 51 32 49 35 
PC254K1 18 14 0 0 0 0 29 41 71 59 
JL24 18 10 0 0 0 33 58 32 42 35 
CG7 16 19 0 0 0 0 59 61 41 39 
Kwarts4 8 5 0 38 37 0 32 32 31 30 
PC287K5 4 8 23 34 0 0 30 21 47 45 
inoculated 
2Uninoculated 
3D. afr/ca/ws-susceptible standard 
4D. afr/ca/ws-tolerant standard 
During 2004-2005 the UBS % of inoculated Harts, Sellie, PC254K1 and JL24 were 
substantially higher than those of their respective uninoculated counterparts. The 
47 
UBS % of all the other genotypes did not differ much between the inoculated and 
uninoculated pairs, although both sections of CG7 were higher than 10 %. Only 
Kwarts and PC287K5 had UBS % lower than 10 % in this trial (Table 3.6). Also in this 
trial the grading from choice to crushing corresponded to the respective UBS % 
(Table 3.6). Choice grade kernels were obtained from the inoculated and the 
uninoculated sections from PC287K5 as well as from the uninoculated Kwarts. 
Standard grade was obtained only from Kwarts in the inoculated section and from 
JL24 and Sellie from the uninoculated section. The rest of the genotypes produced 
only diverse and crushing grade in the inoculated and the uninoculated sections of 
the microplot trial. 
Differences between yield quantity of each genotype in the uninoculated section and 
their corresponding inoculated sections ranged from 81 kg/ha (Kwarts) to 2 170 kg/ha 
(UF85) during 2003-2004 and from 35 kg/ha (JL24) to 1 511 kg/ha (Sellie) during 
2004-2005 (Tables 3.7 & 3.8). Yield quantity was variable over and between 
inoculated and uninoculated sections in both microplot trials (Tables 3.7 & 3.8). 
Table 3.7. Yield and income loss or gain of eight groundnut genotypes from inoculated and 
uninoculated sections of a microplot trial during 2003-2004. 
Yield Yield Yield Income Income 
Genotype (kg/h) loss/gain loss/gain (%) (R/t on) loss/gain (%) 
(Yui) (Yi) (Yui - Yi) (Py1) (lui) (li) (Pin2) 
SellieJ 2 420 2 102 318 -15.1 4 425 2917 -51.7 
PC223 1 645 2 453 -808 + 32.9 4 334 2 864 -51.3 
73-30 2111 1 970 141 -7.2 4212 3 742 -12.6 
UF85 593 2 763 -2 170 + 78.5 3 209 2 889 -11.1 
453 2 699 2 475 224 -9.1 4 073 3 741 -8.9 
Kwarts4 2 653 2 734 -81 + 3.0 5 394 5 242 -2.9 
PC254K1 988 846 142 -16.8 4 750 4 801 + 1.1 
PC287K5 1 752 2 675 -923 + 34.5 3715 4 354 + 14.7 
1Py = (Yui-Yi)100/Yi where Py = yield loss/gain, Yui = yield in uninoculated plots and Yi = 
yield in inoculated plots. 
2Pin = (lui-li)100/li where Pin = income loss/gain, lui = income calculated for uninoculated 
plots and li = income calculated for inoculated plots (Grades were priced according to market 
value at the time of study). 
3 D. africanus- susceptible standard 
4D. a/ncamvs-tolerant standard 
48 
Table 3.8. Yield and income loss or gain of eight groundnut genotypes from inoculated and 
uninoculated sections of a microplot trial during 2004-2005. 
Yield Yield Yield Income Income 
Genotype (kg/h) loss/gain loss/gain (%) (R/ton) loss/gain (%) 
(Yui) (Yi) (Yui - Yi) (Py1) (lui) CO (Pin2) 
Kwarts4 6 036 5 778 258 -4.4 2 513 1 994 -26.0 
Sellie3 5 040 6 551 -1 511 +23.1 1 945 1 710 -13.7 
JL24 5 635 5 600 35 -0.6 1 944 1 777 -9.4 
PC287K5 4 444 4 951 -507 +10.2 2 284 2 094 -9.1 
PC254K1 3 413 2 640 773 -29.3 1 605 1 485 -8.1 
Harts 6 222 6 409 -187 +2.9 1 878 1 855 -1.2 
CG7 5 831 4 382 1 449 -33.1 1 809 1 793 -0.9 
PC299K5 4 160 4 871 -711 +14.6 1 686 1 719 + 1.9 
1Py = (Yui-Yi)100/Yi where Py = yield loss/gain, Yui = yield in uninoculated plots and Yi = 
yield in inoculated plots. 
2Pin = (lui-li)100/li where Pin = income loss/gain, lui = income calculated for uninoculated 
plots and li = income calculated for inoculated plots (Grades were priced according to market 
value at the time of study). 
3D. a/r/canus-susceptible standard 
4D. afr/'canus-tolerant standard 
During 2003-2004 the yield loss/gain % (Py) ranged from a gain of 78.5 % for UF85 
to a loss of 16.8 % for PC254K1 (Table 3.7). Py of the genotypes tested during 2004-
2005 ranged from a gain of 23.1 % for Sellie to a loss of 33.1 % for CG7 (Table 3.8). 
In contrast to the lower yields produced by Sellie, Kwarts, PC254K1 and PC287K5 
during 2003-2004 compared to those produced during 2004-2005, a higher income 
was earned during 2003-2004 compared to that earned during 2004-2005 (Tables 
3.7 & 3.8). A higher income was earned by the genotypes during 2003-2004 because 
the grading prices (R per ton) for each grading class for this season were much 
higher than that of 2004-2005 (Landbouweekblad, 2004 & 2005). The differences in 
income earned for choice grade during 2003-2004 and 2004-2005 were R500 per 
ton, for standard grade R400 per ton and for crushing and diverse grade R300 per 
ton each. Only PC287K5 (14.7 %) and PC254K1 (1.1 %) recorded an increase in 
potential income during 2003-2004 (Table 3.7). During 2004-2005 only PC299K5 
showed an increase in potential income (1.9 %). Compared with the decrease in 
49 
potential income for Sellie (13.7 %) and Kwarts (26 %) during the same season, the 
income of CG7 decreased by 0.9 % and that of 254K1 by 8.1 %. 
3.4.1.2 Field trials 
3.4.1.2.1 Final nematode population densities, reproduction factors and 
reproduction rate 
Nematode populations in the soil from Hartswater and Jan Kempdorp consisted 
mainly of free-living nematodes and the plant-parasitic spp. Meloidogyne and 
Helicotylenchus. At both localities the groundnut genotypes did not differ significantly 
from each other in terms of the number of either Meloidogyne spp. or Helicotylenchus 
spp. per 200 cm3 soil (data not shown). Meloidogyne spp. numbers were low in the 
roots of the genotypes from both sites. The genotypes did not differ significantly from 
each other in terms of root-knot nematode numbers per 50 g roots (data not shown). 
D. africanus was present in the roots of the groundnut genotypes from the Hartswater 
trial (Fig. 3.4), but it did not occur in roots of the genotypes from the Jan Kempdorp 
trial. 
8.6 
6.6 
o 
2 4.6 
en 
ir> 
l 26 
0.6 
-1.4 
Figure 3.4. Ditylenchus africanus numbers (Pf) in the roots of eight groundnut 
genotypes from a field trial planted at Hartswater during 2004-2005 
(P < 0.05; F-ratio = 4.43). 
50 
LSD (P< 0.05) 
O 
O Q. O Q. 
Genotype 
D. africanus numbers in the roots of CG7 and PC254K1 from Hartswater did not 
differ significantly from those in the roots of Sellie and JL24, but were significantly 
lower than those in roots of the other genotypes (Fig. 3.4). 
Significantly lower D. africanus numbers were extracted from CG7 and PC254K1 
pods than from those of the rest of the genotypes planted at both Hartswater and Jan 
Kempdorp (Figs 3.5 & 3.6). At Jan Kempdorp D. africanus numbers of PC287K5 
were significantly higher than those in CG7 and PC254K1 but were significantly lower 
than those in pods of the rest of the genotypes (Fig. 3.6). Pf of D. africanus in pods of 
CG7 was 347.3-fold lower than that of Sellie at Hartswater and 142-fold lower than 
that of Sellie at Jan Kempdorp. In pods of PC254K1 the Pf was 9.3-fold lower than 
that of Sellie at Hartswater and 142-fold lower than that of Sellie at Jan Kempdorp. 
7.5 -
5.5 
i 3.5 
1.5 
-0.5 
I I I I I LSD (P_< 0.05) I 
I 
I 
I *: * 
O h-
O CO 
CM CM 
O O 
Q. Q. 
Genotypes 
Figure 3.5. Ditylenchus africanus numbers (Pf) in pods of eight groundnut genotypes 
from a field trial planted at Hartswater during 2004-2005 (P < 0.05; F-
ratio = 15.38). 
51 
5.5 
I 
S. 3.5 
-0.5 
-
LSD (P< 0.05) 
: I I I I I 
- I 
- I I 
^ ^ CO a 
O o D_ D_ 
o 
Genotype 
Figure 3.6. Ditylenchus africanus numbers (Pf) in pods of eight groundnut 
genotypes from a field trial planted at Jan Kempdorp during 
2004-2005 (P < 0.05; F-ratio = 44.06). 
The numbers of D. africanus in pods of Sellie was used as the standard to determine 
the RR rate of the nematode in pods of the other genotypes. The RR of CG7 was 0 
% at both Hartswater and Jan Kempdorp (Table 3.9). 
Table 3.9. Ditylenchus africanus numbers (Pf) and reproduction rates (RR) in pods of eight 
groundnut genotypes from field trials planted at Hartswater and Jan Kempdorp 
during 2004-2005. 
Genotype Pf (15 Hartswater g pods) Jan Kempdorp Hartswater RR' Jan Kempdorp 
Sellie2 1 042 142 100 100 
Kwarts3 851 150 81.6 105.6 
PC299K5 805 148 77.3 104.2 
PC287K5 785 52 75.3 36.6 
Harts 371 180 35.6 126.8 
JL24 328 105 31.5 73.9 
PC254K1 112 0 10.8 0 
CG7 3 0 0 0 
1RR = Pf (genotype) / Pf (Sellie) x 100 
2D. afr/cantvs-susceptible standard 
3D. afr/cantvs-tolerant standard 
52 
RR of PC254K1 was 0 % at Jan Kempdorp and 10.8 % at Hartswater. For PC287K5 
the RR was 75.3 % at Hartswater and 36.6 % at Jan Kempdorp. 
3.4.1.2.2 Yield assessments 
At Hartswater (Table 3.10) the UBS % of Harts was 39 % and that of Sellie was 12 
%. The yields of these latter two genotypes were downgraded to diverse and 
crushing grade. For the rest of the genotypes the UBS % was lower than 10 %. CG7 
(4 %), PC287K5 (3 %) and PC254K1 (4 %) had the lowest UBS % and they also 
produced choice grade kernels. The rest of the genotypes, including the tolerant 
Kwarts produced standard, diverse or crushing grade. 
At Jan Kempdorp (Table 3.11) the UBS % of Harts was 23 % and it was downgraded 
to diverse and crushing grade. The UBS % of the rest of the genotypes was all below 
10 % and all produced choice grade kernels. 
Table 3.10. Yield quality of eight groundnut genotypes from a field trial planted at Hartswater 
during 2004-2005. 
Genotype UBS 
(%) 
Choice 
(%) 
Standard 
(%) 
Diverse 
(%) 
Crushing 
(%) 
Harts 39 0 0 46 54 
Sellie1 12 0 0 79 21 
Kwarts2 9 0 53 28 19 
JL24 8 0 59 24 17 
PC299K5 7 0 50 35 15 
CG7 4 61 0 25 14 
PC254K1 4 59 0 27 14 
PC287K5 3 31 0 33 36 
1D. afr/cantvs-susceptible standard 
2D. a/r/cantvs-tolerant standard 
53 
Table 3.11. Yield quality of eight groundnut genotypes from a field trial planted at Jan 
Kempdorp during 2004-2005. 
Genotype UBS 
(%) 
Choice 
(%) 
Standard 
(%) 
Diverse 
(%) 
Crushing 
(%) 
Harts 23 0 0 81 19 
PC287K5 4 62 0 23 15 
Kwarts1 3 74 0 20 6 
JL24 3 61 0 31 8 
PC299K5 3 55 0 33 12 
CG7 3 50 0 36 14 
Sellie2 1 59 0 34 7 
PC254K1 0 63 0 30 7 
D. a/r/'canus-tolerant standard 
2D. afr/canus-susceptible standard 
3.4.2 Reproduction and damage threshold-levels of D. africanus on resistant, 
susceptible and tolerant groundnut genotypes 
3.4.2.1 Final nematode population densities and reproduction factor 
Pf in pods of PC254K1 did not always increase with an increase in Pi but the Pf of 
PC287K5 did (Table 3.12). Pf of Sellie and Kwarts also increased with an increase in 
Pi, except at Pi = 5 000 and higher for Sellie and at Pi = 2 500 and higher for Kwarts. 
The RF of Sellie and Kwarts was higher than one at all but the highest Pi's, viz. from 
Pi = 5 000 for Kwarts and Pi = 10 000 for Sellie (Table 3.12). The RF of PC254K1 
and PC287K5, however, remained lower than one at all corresponding Pi's. 
54 
Table 3.12. Final population densities (Pf) and reproduction factor (RF) of Ditylenchus 
africanus in pods of four groundnut genotypes inoculated at planting with 
escalating initial population densities (Pi) in a microplot trial during 2004-2005. 
Pi Pf(15gpods) RF
1 
Sellie2 Kwarts3 PC254K1 PC287K5 Sellie Kwarts PC254K1 PC287K5 
0 0 0 0 0 - - - -
50 377 856 16 12 7.54 17.12 0.32 0.24 
250 450 366 38 0 1.80 1.46 0.15 0 
1 000 2 361 1 228 11 13 2.36 1.23 0.01 0.01 
2 500 4 462 2 562 32 58 1.78 1.03 0.01 0.02 
5 000 9 586 1 535 83 234 1.91 0.31 0.02 0.05 
10 000 6 261 6 146 574 423 0.63 0.62 0.06 0.04 
20 000 8 628 4 332 405 671 0.43 0.22 0.02 0.03 
1RF = Pf/Pi 
2D. a/h'canus-susceptible standard 
3D. afr/canus-tolerant standard 
The relationships between Pi and Pf were best described by non-linear equations for 
Sellie, Kwarts, PC254K1 and PC287K5 (Fig. 3.7). 
55 
4.0i 
3.5 
+ x 
Sellie y = 3.826 -2.837 (0.99792 A X) 
RA2 =0.70 
kwarts y = 3.646 -0.999 (0.999615 AX) 
RA2 =0.74 
3.0"| / pC287K5 y = 2.781 -2.459 (0.999653 A X) 
RA2 =0.85 
2.5- " 
2.0" 
1.5-
1.0" 
0.5-
/ i 
PC254K1 y = 2.679 -0.00182 * X) (0.999433 A X) 
RA2 =0.64 
o o 
in 
O O 
O 
O o 
in 
CM lO r~-
o o 
o 
o 
o o o o 
o 
in 
o o o o 
o o 
CM 
Pi 
Figure 3.7. Non-linear relationships between initial nematode population densities 
(Pi) and final nematode population densities (Pf) in pods of Sellie, 
Kwarts, PC254K1 and PC287K5 in a microplot trial during 2004-2005. 
R2 values were significant for all the genotypes. Exponential models explained 70 % 
of the variation for Sellie, 85 % for PC287K5, 74 % for Kwarts and 64 % for 
PC254K1. The regression lines levelled off from Pi = 2 500 for Sellie and Pi = 5 000 
for Kwarts but at Pi = 7 500 for PC254K1 and PC287K5. 
3.4.2.2 Yield assessments 
The UBS % of Sellie was substantially higher than those of the D. afr/canus-tolerant 
Kwarts and the two breeding lines (Table 3.13). PC287K5 had the lowest overall 
UBS %. Income from the four genotypes varied over the different Pi levels (Table 
3.13). 
56 
Table 3.13. Yield quality and income for four groundnut genotypes inoculated at planting with 
a range of initial Ditylenchus africanus population densities (Pi) in a microplot trial 
during 2004-2005. 
Genotype Pi UBS 
(%) 
Choice 
(%) 
Standard 
(%) 
Diverse 
(%) 
Crushing 
(%) 
Income 
(R/ha) 
Sellie' 0 2 27 0 40 33 4 313 
50 5 0 35 38 27 4 879 
250 6 0 32 40 28 4 178 
1 000 6 0 26 43 31 3 638 
2 500 13 0 0 63 37 3 357 
5 000 20 0 0 65 35 4 184 
10 000 22 0 0 50 50 2 854 
20 000 26 0 0 45 55 2 242 
Kwarts2 0 3 0 28 38 34 2 807 
50 1 38 0 39 23 5 485 
250 1 31 0 41 28 4 921 
1 000 3 0 34 45 21 4510 
2 500 5 0 33 45 22 4 194 
5 000 5 0 28 47 25 4 298 
10 000 6 0 26 48 26 3 608 
20 000 11 0 0 65 35 3 292 
PC254K1 0 4 0 19 31 50 4 348 
50 2 0 15 30 55 3 980 
250 4 25 0 27 48 3 224 
1 000 2 26 0 33 41 3 335 
2 500 5 0 0 50 50 3 288 
5 000 3 0 20 35 45 4 183 
10 000 3 0 12 20 68 3 362 
20 000 3 0 17 20 63 2 796 
PC287K5 0 0.4 34 0 34 32 3 874 
50 0.7 27 0 36 37 4 399 
250 1 40 0 34 28 4 608 
1 000 0.6 26 0 29 45 2 431 
2 500 0.1 32 0 34 34 3 636 
5 000 2 25 0 37 38 3 052 
10 000 1 23 0 35 42 2 874 
20 000 3 0 29 36 35 2 780 
1D. afr/canus-susceptible standard 
2D. afr/canus-tolerant standard 
57 
The relationships between Pi and UBS % in this trial were best described with a non 
linear regression for Sellie (exponential) and linear regressions for PC287K5 and 
Kwarts. The variation of UBS % for PC254K1 at the different Pfs was not significant 
and could, therefore, not be fitted to any model. R2 values were significant for the 
three former genotypes and the models explained 97 % of the variation for Sellie, 91 
% for Kwarts and 63 % for PC287K5 (Fig. 3.8). 
30 
25 
20 
& 15 m 
10 
Sellie y = 26.07 -22.77 (0.9997864 A X) 
RA2 = 0.97 .,-----
Kwarts y = 0.0004090 X + 2.562 
RA2 = 0.91 
?4 ... 
"" o PC287K5 y = 0.0001513 X + 0.852 
0 RA2 = 0.63 
o o o o o o o o o o o o o o o 
in o in o in o o 
CM m h- o C\J in h- o 
CM 
Pi 
Figure 3.8. Relationships between initial nematode population 
densities (Pi) and UBS % of Sellie, Kwarts, PC254K1 
and PC287K5 in a microplot trial during 2004-2005. 
The yield quantities of none of the genotypes corresponded with an increasing Pi 
(Table 3.14). 
58 
Table 3.14. Yield of four groundnut genotypes inoculated at planting with a range of initial 
Ditylenchus africanus population densities (Pi) in a microplot trial during 2004-
2005. 
Pi Yield (kg / ha) Sellie1 Kwarts^ PC254K1 PC287K5 
0 1 863 1 446 2 467 1 597 
50 2 390 2116 2 348 1 952 
250 2219 2 030 1 523 1 820 
1 000 1 842 2 155 1 505 1 119 
2 500 1 835 2 019 1 934 1 540 
5 000 2 263 2114 2 322 1 377 
10 000 1 680 1 792 1 872 1 337 
20 000 1 356 1 779 1 505 1 442 
D. afr/canus-susceptible standard 
2D. africanus-tolerant standard 
3.4.3 Expression of D. africanus resistance in callus tissue 
3.4.3.1 Final nematode population densities and reproduction factor 
There were no significant differences between the nematode numbers in Sellie, 
Kwarts, PC287K5 and PC254K1 callus tissue after four weeks' incubation although 
D. africanus increased from 224- to 393-fold on average over this period (Table 
3.12). The RF of D. africanus was much higher than one on all callus tissue of the 
four groundnut genotypes (Table 3.12). 
59 
Table 3.15. Ditylenchus africanus numbers (Pf) and reproduction factors (RF) on callus tissue 
of four groundnut genotypes incubated four weeks at 28 °C in a growth cabinet 
trial during 2008. 
Genotype Pf (1 g callus) RF1 
Sellie 3 355 335.5 
Kwarts 3 928 392.8 
PC287K5 2 240 224.0 
PC254K1 2 728 272.8 
1RF = Pf/Pi 
3.5 DISCUSSION 
Contrary to the high Pf of D. africanus in the susceptible standard Sellie and most of 
the other genotypes tested during the identification of resistance part of this study, 
PC254K1 and CG7 pods consistently maintained significantly lower Pf s at harvesting 
under microplot as well as field conditions. This is fair proof of their resistance (Bos & 
Parlevliet, 1995) to D. africanus. Low Pf in PC287K5 in the microplots also indicated 
resistance but high Pf in the field shows that the resistance in this line may be lower 
or less sustainable under field conditions. The superior resistance of PC254K1 and 
CG7 to D. africanus was also evident in the RF because it generally remained below 
one compared to those of the susceptible standards (Roberts & May, 1986; Windham 
& Williams, 1988). It was further substantiated by the corresponding RR values that 
remained below 10 % (Abdel-Momen et a/., 1998; Hussey & Janssen, 2002; Timper 
et a/., 2003) for these two lines under microplot as well as field conditions. The field 
trials in this part of the study provided solid confirmation of the resistance or 
susceptibility levels of the genotypes tested (De Waele et a/., 1989; Venter et a/., 
1992; Mc Donald, 1998). Although the presence of multiple plant-parasitic nematode 
species in a field may affect the expression of resistance (Barker & Olthof, 1976; 
Eisenback, 1985) this did not seem to be applicable to D. africanus resistance in 
PC254K1 and CG7 as indicated in this part of the study. 
Increase or decrease in yield quality of the genotypes in all the microplot trials of this 
part of the study did not correlate with Pf and confirm similar results from previous 
studies (Venter et a/., 1991 & 1992; Mc Donald et al., 2005). Therefore, the much 
higher yield recorded for UF85 (2003-2004) and Sellie (2004-2005) in the inoculated 
section of the respective microplot trials compared to that of the uninoculated 
60 
sections is difficult to explain. It might in part be due to conjugating environmental 
factors (Niblack et al., 1986a), which did not seem to be consistent, however. 
The higher UBS % of most genotypes in the inoculated sections of the microplot trials 
compared to the uninoculated sections agrees with several other studies in that the 
main effect of D. africanus on groundnut yield is qualitative (Jones & De Waele, 1988 
& 1990; De Waele et al., 1989; Venter et al., 1991 & 1993; Mc Donald et al., 2005). 
In the presence of aggressive nematodes environmental factors affecting kernel 
quality (Barker & Noe, 1987; Mc Donald et al., 2005) often play a lesser role in 
symptom expression (Barker & Noe, 1987). This may also be applicable to D. 
africanus since UBS % of yields obtained from inoculated sections was generally 
higher compared to the uninoculated counterparts of most genotypes. However, the 
generally lower UBS % of PC254K1 and CG7 yields compared to those of the other 
genotypes supports trends in Pf, RF and RR data and confirms the resistance 
particularly of these two genotypes to D. africanus under microplot as well as field 
conditions. 
The superior resistance of PC254K1 to D. africanus in the microplot trial on the 
reproduction and damage potential part of this study was confirmed because Pf did 
not increase as a function of increasing Pi and remained low even at the highest Pi's 
(Bos & Parlevliet, 1995). Intra-species competition (Seinhorst, 1965) may have been 
triggered in PC287K5 when Pi reached 2 500 nematodes per plant and higher but 
this did not occur in PC254K1 because this genotype did not support reproduction of 
this nematode even at the lowest Pi levels. Nematode multiplication-threshold levels 
(Cook & Starr, 2006) were probably reached at Pi = 5 000 and higher for Sellie and 
at Pi = 2 500 and higher for Kwarts. At these levels the D. africanus population 
increased to such an extent in the pods of the latter two genotypes that resources for 
further growth and reproduction of the parasite were depleted (Cook & Starr, 2006). 
Above this multiplication threshold level the Pf of parasites also becomes less than 
the initial density in a susceptible host (Cook & Starr, 2006). A D. africanus 
multiplication threshold level was not reached in PC287K5 or PC254K1 during this 
part of the study because the Pf in pods of the latter two genotypes never exceeded 
the corresponding Pi (Cook & Starr, 2006). High Pf in pods of Sellie and Kwarts at 
lower Pi's as in previous studies (Basson et al., 1993; Mc Donald et al., 2005) 
showed that survival of D. africanus in either small or large numbers at the beginning 
of a growing season is irrelevant because of the nematode's high reproduction 
potential (De Waele & Wilken, 1990). The resistance of PC254K1 to D. africanus was 
61 
furthermore substantiated by the RF, which remained below one (Roberts & May, 
1986; Windham & Williams, 1988) at all Pi levels. Based on RF > 1 the nematode 
was able to reproduce sufficiently on Sellie and Kwarts at lower Pi levels, however 
(Windham & Williams, 1988). The non-linear relationship between Pf and UBS % for 
Seilie in the trial on reproduction and damage-threshold levels is similar to that 
observed previously in microplot studies, which propose that UBS % is a reliable 
estimate of the effect of this nematode (Mc Donald et al., 2005). Compared with 
Sellie and Kwarts the resistance of PC254K1 to D. africanus is, therefore, also 
confirmed by its high yield quality, having produced only choice and standard grade 
kernels at all Pi levels. 
The high Pf and RF of D. africanus on callus tissue of Sellie, Kwarts, PC287K5 as 
well as on those of PC254K1 confirms that D. africanus is able to reproduce optimally 
on callus tissue of all four groundnut genotypes (Windham & Williams, 1988). These 
results support previous studies, which indicated that callus tissue of plants resistant 
to a specific nematode in nature will often support good reproduction of that 
nematode on callus tissue of the same resistant plant (Webster & Lowe, 1966; 
Krusberg & Babineau, 1977). The higher reproduction rate of nematodes on callus 
tissues compared to that on resistant hosts has previously been reported (Krusberg 
& Babineau, 1977). According to Webster and Lowe (1966), plant growth-regulating 
substances such as 2.4-D used in the medium on which the callus tissue was 
cultured during this study (Van der Walt & De Waele, 1989) may have played a role 
in making incompatible cells compatible to nematodes in callus tissue (Webster & 
Lowe, 1966). Nematode resistance of groundnut genotypes can, therefore, not be 
evaluated against D. africanus based on Pf and RR obtained from their callus tissue. 
This part of the study did, however, confirm the high reproduction rate of D. africanus 
in in vitro callus tissue from groundnut leaves that was previously reported by Van 
der Walt and De Waele (1989) in their studies. 
This is the first report of groundnut genotypes that express high-level, sustainable 
resistance to D. africanus under microplot as well as field conditions. PC254K1 will 
play an important role in groundnut-breeding programmes because of its evidently 
high level of resistance, even at high nematode infestation levels. Although this line 
may lack many desirable traits required for an agronomically acceptable cultivar e.g. 
high yield potential, desired kernel size, colour and form it should be acceptable for 
use to introgress resistance into preferred breeding material. A high-yielding cultivar 
62 
developed from PC254K1 should also be able to produce better kernel quality, which 
may increase the net income per ha (Mc Donald et a/., 2005). 
Although only tested in one part of this study, there are strong indications that CG7 
may also have superior resistance to D. africanus. It warrants further investigation 
should a comprehensive programme on the introgression of nematode resistance in 
groundnut be initiated. 
Although PC287K5 also maintained low nematode numbers in some trials, its level of 
resistance does not seem to be as high as that of PC254K1. However, it still 
performed well compared to tolerant cultivar Kwarts, which might indicate that this 
line could also be tolerant. Therefore, it may still play an important role, particularly 
under lower nematode population pressure. 
63 
CHAPTER 4 
REPRODUCTIVE AND DAMAGE POTENTIAL OF FIVE GEOGRAPHICALLY-
ISOLATED DITYLENCHUS AFRICANUS POPULATIONS ON GROUNDNUT 
4.1 INTRODUCTION 
From an economic and environmental perspective host-plant resistance is one of the 
most preferred management tools for plant-parasitic nematodes on a variety of crops 
(Starr et al., 2002; Agudelo et al., 2005; Dickson & De Waele, 2005; Cook & Starr, 
2006). This also applies to the management of D. africanus on groundnut (De Waele 
et al., 1990). The breeding line PC254K1 was confirmed to be highly resistant to D. 
africanus (Chapter 3) and showed potential for inclusion in the local groundnut-
breeding programme as a primary source of resistance. 
Acceptance of a new cultivar developed from PC254K1 on the market will not only 
depend on the agronomic acceptability of such a cultivar, but also on the 
sustainability of its resistance to D. africanus. Sustainability of resistance may in turn 
depend on the virulence and damage potential of nematode populations present in 
soils where the crop is produced (Blok et al., 1997; Thies & Ferry, 2002). Potential 
differences in reproduction and damage potential of geographically-isolated D. 
africanus populations will, furthermore, affect the composition of a suitable 
management system for a specific production area (Sree Latha et al., 1989; Noe, 
1992). Obviously the usefulness of a specific resistant cultivar for each area is also 
dependent on the damage potential of a specific plant-parasitic nematode for a 
particular area. Knowledge of any variability between and within D. africanus 
populations present in the various groundnut production areas is, therefore, 
necessary for development of cultivars of which their resistance will be sustainable 
for all respective areas of production (Agudelo et al., 2005). 
Although a number of studies have been done on the reproduction of D. africanus on 
groundnut callus tissue (Van der Walt & De Waele, 1989) at temperatures that 
ranged from 16 °C to 34 °C (De Waele & Wilken, 1990) and on the reproduction and 
damage potential of D. africanus on a number of groundnut genotypes (Basson et al., 
1991, 1992 & 1993; Venter et al., 1991 & 1993; Van der Merwe & Joubert, 1994; Mc 
Donald et al., 2005), no information is available on the comparative reproduction and 
damage potentials of D. africanus populations isolated from different geographical 
64 
locations. The purpose of this study was to establish under controlled and semi-
controlled conditions whether there are differences in the reproduction and damage 
potential of D. africanus originating from different localities in the groundnut-
producing areas of South Africa. 
4.2 MATERIALS AND METHODS 
4.2.1 D, africanus populations 
Callus used for propogation of the various geographically-isolated D. africanus 
populations was initiated from young, surface-sterilised leaves of Sellie (Chapter 2). 
D. africanus populations used for the inoculation of callus tissue were extracted from 
infected groundnut pods that were collected from infested fields from Mareetsane 
(26.15°S, 25.43°E) (population 1), Jan Kempdorp (27.95°S, 24.85°E) (population 2), 
Vaalharts (27.83°S, 24.79°E) (population 3), Schweizer-Renecke (27.19°S, 25.33'E) 
(population 4) and Theunissen (28.40°S, 26.71°E) (population 5) (Fig. 4.1). 
Figure 4.1. Diiylenchus africanus populations were isolated from infected groundnut pods 
collected from five different localities (indicated by red dots) within the 
groundnut-production area of South Africa. 
65 
These infected pods were collected from groundnut shelling plants at the 
abovementioned locations by officers of the Perishable Products Export Control 
Board (PPECB) of South Africa. The material was delivered to the Nematology Unit 
of ARC-GCI in Potchefstroom within two to three days of collection. After collection 
the pods were put in dry cooler bags and were stored at room temperature under air-
dry conditions until extraction of the nematodes. Twenty pods from each locality were 
shelled by hand and a mean number of 251 484 D. africanus was extracted from the 
pod tissues (5 g hull and 5 g kernel) from each locality using the soaking method 
(Chapter 2). In vitro, aseptic calluses were then established of each of the five above 
mentioned D. africanus populations with five males and five females picked from the 
extracted D. africanus pod-samples following procedures described in Chapter 2. 
4.2.2 Trial layout 
4.2.2.1 Growth cabinet trial 
This trial was conducted during July / August 2007. Three growth cabinets were set 
at 21 °C, 28 °C and 35 °C, respectively, and allowed to stabilise for three days at the 
respective temperature before proceeding with this trial. Ninety calluses of + 1.48 g 
each were used. Nematodes inoculated on the callus were extracted from cultures 
established for each locality (4.2.1). Each of the ninety calluses was inoculated with 
five males and five females of the respective D. africanus populations following the 
relevant procedures described in Chapter 2. 
The trial was laid out in randomised complete block, factorial design. The three 
growth cabinets with the different temperature regimes were the main factor. The 
sub-factor consisted of callus tissue cultures of each of the five D. africanus 
populations (five treatments), each replicated six times. The inoculated calluses were 
incubated in darkness for four weeks in the growth cabinets to allow the nematodes 
to complete a minimum of four life cycles at a specific temperature regime (De Waele 
& Wilken, 1990). After this the nematodes were extracted from the callus tissue 
according to the procedures described in Chapter 2. The nematodes from each 
callus were counted under a research microscope and the nematode (Pf) numbers 
were expressed as D. africanus per 1 g callus. 
66 
4.2.2.2 Greenhouse trial 
This trial was conducted between May and October 2007. Sixty plastic pots were 
filled with 4 000 cm3 sandy-loam soil fumigated with EDB six weeks before planting 
(Chapter 2). Nutrients were added and mixed into the soil in each pot as desribed in 
Chapter 2. Nutrient requirements for this trial were 5.84 g dolomitic lime, 3.52 g super 
phosphate (10.5 % P), 2.32 g sodium chloride (KCI) and 1.50 g calsium nitrate 
(CaN03) per pot. One seed was planted to a depth of 5 cm per pot (Chapter 2). A 
plastic marker per pot indicated the genotype and replicate numbers. Sellie and 
PC254K1 seeds were obtained from nematode-free mircoplot nurseries (Chapter 2) 
and treated with fungicide and nitrogen-fixing bacteria as described in Chapter 2. 
The trial was planted in a randomised complete split-plot design in a greenhouse with 
a temperature regime of 18 °C to 27 °C with a 13-h photoperiod. Factor one was a 
plot of 30 pots each of Sellie (suscepitble) or PC254K1 (resistant). The trial included 
two factors and six single-plant replicates inoculated with each of the five D. 
africanus populations. Each pot was inoculated at planting with + 2 000 D. africanus 
consisting of various life stages. The nematodes used for inoculation were extracted 
from in vitro cultures of the five D. africanus populations (4.2.1) and prepared for 
inoculation according to procedures described in Chapter 2. A plastic marker per pot 
indicated the genotype, D. africanus population and replicate number. Pots were 
rotated clockwise every fortnight to limit the effects of temperature or air-flow 
gradients in the greenhouse. Pots were watered three times a week by hand. 
Nematode assessments were made at harvesting on all the plants. D. africanus was 
extracted from pegs, hulls and kernels following the respective methods described in 
Chapter 2. Nematodes were counted under a research microscope and expressed as 
Pf per 15 g pods (Chapter 2). Yield assessments were not made from the single-
plant pots in the greenhouse trial because the technique used for nematode 
extraction is destructive (Bolton et al., 1990) and the amount of kernels left for 
reliable yield assessments was insufficient. 
4.2.2.3 Microplot trial 
This trial was conducted during 2007-2008 in concrete-pipe microplots (Chapter 2). 
All sixty plots were filled with the same fumigated sandy-loam soil used in the 
greenhouse trial (4.2.2.2). Nutrients incorporated into the soil of each plot were 70 g 
67 
dolomitic lime, 51 g super phosphate (10.5 % P), 34 g KCI and 20 g CaN03. Each 
plot was then planted with six seeds respectively of Sellie or PC254K1 to a depth of 5 
cm (Chapter 2). Seeds of the two genotypes were obtained from nematode-free 
nurseries, treated with fungicide and inoculated with Bradyrhizobium arachis before 
planting as described in Chapter 2. 
The trial was planted in a randomised complete split-plot design similar to that of the 
greenhouse trial (4.2.2.2) with two factors, five treatments and six replicates each. 
Factor one consisted of 30 plots of six plants each of Sellie and 30 plots of six plants 
each of PC254K1. The inoculum density was + 2 000 D. africanus of various life 
stages per plant at planting. D. africanus used for inoculation of the plants were 
extracted from the same in vitro cultures of the five geographically-isolated D. 
africanus populations (factor two: treatments) used in the greenhouse trial (4.2.2.2). 
Every treatment was replicated six times. Preparation of D. africanus inoculum and 
inoculation of each plant was performed as described in Chapter 2. A plastic marker 
indicated the genotype, D. africanus population and the replicate number of each 
plot. Supplementary to rainfall plots were irrigated three times a week through a 
micro-sprayer placed in the centre of each plot (Chapter 2). 
Nematode and damage assessments were done at harvesting. Separate nematode 
extractions were done from peg, hull and kernel samples collected from two 
randomly picked plants from each plot. D. africanus was extracted following the 
methods described in Chapter 2. Nematodes were counted under a research 
microscope and expressed as Pf per 15 g pods (Chapter 2). 
The four remaining plants in each plot not used for nematode extraction were used 
for damage assessments. Yield quantity was not determined for this part of the study 
because D. africanus does not affect yield quantity (Chapter 3; Venter et al., 1991 & 
1992; Mc Donald et al., 2005). Yield quality was determined by following standard 
grading procedures (Chapter 2). 
4.3 STATISTICAL ANALYSES 
Unless otherwise stated nematode data were ln(x+1) transformed before being 
subjected to a factorial analysis of variance (ANOVA) (Stat Graphics 5 Plus for 
Windows). Means were separated by a LSD test (P < 0.05). 
68 
Nematode variables included final population densities (Pf) of D. africanus present in 
callus tissue cultures in the growth cabinet trial and Pf present in pods of Sellie and 
PC254K1 in the greenhouse and microplot trials. The nematode reproduction factor 
(RF) in callus tissue cultures (growth cabinet trial) and pods of Sellie and PC254K1 
(greenhouse and microplot trials) were determined as described in Chapter 2. 
4.4 RESULTS 
4.4.1 Growth cabinet trial 
4.4.1.1 Final nematode population densities and reproduction factor 
There was no significant interaction between temperature regime and D. africanus 
population numbers. However, there were significant differences between the 
populations at 21 °C as well as at 28 °C (Fig. 4.2). In the 21-°C regime the Pf of 
population 4 (Schweizer-Renecke) was significantly higher than that of population 1 
(Mareetsane), 2 (Jan Kempdorp) and 5 (Theunissen) (Fig. 4.2). 
10 LSD (P< 0.05} 
8 '? I T l I 
+ :- II 
iff 
8 A -
5 
2 
0 
r -*" 
28 C 
35 C 
D. africanus population 
Figure 4.2. Final D. africanus numbers (Pf) on groundnut callus tissue inoculated with 
five geographically-isolated Ditylenchus africanus populations from Mareetsane 
(population 1), Jan Kempdorp (population 2), Vaalharts (population 3), 
Schweizer-Renecke (population 4) and Theunissen (population 5) incubated for 
four weeks at 21 °C, 28 °C or 35 °C in three separate growth cabinets during 
2007 (P < 0.05; F-ratio = 4.21). 
69 
Populations 1, 2, 3 (Vaalharts) and 5 did not differ significantly from each other. In 
the 28-°C regime the Pf of population 4 was significantly higher than that of 
populations 2 and 3. No significant differences existed between the Pf of the five 
populations at the 35-°C regime. 
Significant differences existed in terms of the Pf of the five populations when it was 
pooled over each of the three temperature regimes (Fig. 4.3). Pf of the populations 
pooled over 28 °C were significantly higher than those pooled over 21 °C and 35 °C. 
However, Pf of the populations on callus tissue pooled over 21 °C and 35 °C did not 
differ significantly from each other. 
-
I LSD (P< 0.05) 
H I 
-
21 28 35 
Temperature regime ("C) 
Figure 4.3. Final D. africanus numbers (Pf) on groundnut callus tissue inoculated 
with five geographically-isolated Ditylenchus africanus populations 
from Mareetsane (population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4) 
and Theunissen (population 5) pooled over three temperature 
regimes, respectively, in a growth cabinet trial during 2007 
(P < 0.05; F-ratio = 76.69). 
The RF of all five populations at all three temperature regimes was higher than one 
(Table 4.1). The RF of the populations at 28 °C were + 58- to 373-fold higher than 
those of the same populations on callus tissues at 21 °C and + 35- to 156- fold higher 
than those of the same populations at 35 °C. The RF rates at 35 °C were on average 
only slightly higher than those at 21 °C. 
70 
Table 4.1. Reproduction factor (RF) of five geographically-isolated Ditylenchus afhcanus 
populations from Mareetsane {population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4) and Theunissen 
(population 5) on groundnut callus incubated four weeks at 21 °C, 28 0C and 35 
°C in separate growth cabinets during 2007. 
Nematode RF1 
population 21 °C 28 °C 35 °C 
11.0 1 286.7 8.6 
3.4 650.8 5.1 
3.0 174.0 4.6 
2.3 856.8 5.5 
1.5 126.3 3.6 
4.2 587.5 5.5 
1RF = Pf/Pi 
4.4.2 Greenhouse trial 
4.4.2.1 Final nematode population densities and reproduction factor 
A significant interaction existed between the groundnut genotypes and the Pf of the 
five D. afhcanus populations in the greenhouse trial. Pf of all five populations was 
significantly higher in Sellie than in PC254K1 (Fig. 4.4). No significant differences 
existed between the five populations on Sellie. There were significant differences in 
Pf of the populations in PC254K1, however (Fig. 4.4). Pf of population 2 (Jan 
Kempdorp) was significantly lower than those of the rest. 
71 
10 
4 -
- LSD (P < 0.05) 
[ I
 1
 1
 1 (-H 1 I I I 
- I I I I 
-
I 
Sellie 
PC254K1 
12 3 4 5 
D. africanus population 
Figure 4.4. Final Ditylenchus africanus numbers (Pf) in pods of Sellie and PC254K1 
inoculated at planting with five geographically-isolated Ditylenchus africanus 
populations from Mareetsane (population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4) and Theunissen 
(population 5) in a greenhouse trial during 2007 (P < 0.05; F-ratio = 2.92). 
The RF of all five populations on Sellie were higher than one (Table 4.2). In contrast 
to this the RF on PC254K1 were all lower than one. The reproduction of the five 
populations was on average + 51-fold lower on PC254K1 than on Sellie. 
Table 4.2. Reproduction factors (Pf) of five geographically-isolated Ditylenchus africanus 
populations from Mareetsane (population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4} and Theunissen 
(population 5) in pods of Sellie and PC254K1 in a greenhouse trial during 2007. 
Nematode RF1 
population Sellie PC254K1 
2 3.94 0.02 
4 3.82 0.10 
1 3.59 0.09 
5 3.25 0.11 
3 3.22 0.05 
Mean 3.56 0.07 
1RF = Pf/Pi 
72 
4.4.3 Microplot trial 
4.4.3.1 Final nematode population densities and reproduction factor 
No significant interaction existed between the five populations and the two groundnut 
genotypes. Pf of the populations on Sellie and PC254K1 were relatively low for all 
five populations (Fig. 4.5). Although the Pf of all of the corresponding nematode 
populations were higher in Sellie than in PC254K1 there were no significant 
differences between the respective populations in either genotype. 
- LSD (P< 0.05) 
[ I i i i I 
; i i i i I 
12 3 4 5 
D. africanus population 
Figure 4.5. Final Ditylenchus africanus numbers (Pf) in pods of Sellie and PC254K1 
inoculated at planting with five geographically-isolated Ditylenchus africanus 
populations from Mareetsane (population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4) and Theunissen 
(population 5) in a microplot trial during 2007-2008 (P < 0.05; F-ratio = 75.76). 
The low Pf of the five populations in the pods of Sellie and PC254K1 were also 
reflected in their respective low RF, which remained lower than one for both 
genotypes (Table 4.3). 
73 
Table 4.3. Reproduction factor (Pf) of five geographically-isolated Ditylenchus africanus 
populations from Mareetsane (population 1), Jan Kempdorp (population 2), 
Vaalharts (population 3), Schweizer-Renecke (population 4) and Theunissen 
(population 5) in pods of Sellie and PC254K1 in a microplot trial during 2007-
2008. 
Nematode RF2 
population Sellie PC254K1 
4 068 O03 
1 0.51 0.08 
3 0.29 0.02 
2 0.27 0.03 
5 0.25 0.02 
Mean 0.40 0.04 
1RF = Pf/Pi 
In spite of the low Pf, however, the average reproduction on PC254K1 was still only 
one tenth of that of Sellie. 
4.4.3.2 Damage assessments 
UBS % of PC254K1 varied and ranged from 4 % (population 3) to 9 % (populations 4 
and 5) (Table 4.4). PC254K1 produced choice grade (population 3) or standard 
grade (populations 1, 2, 4 and 5). In contrast with the UBS % of PC254K1 kernels 
those of Sellie were all higher than 10 %, ranging from 12 % to 14 %. Sellie kernels 
graded crushing and diverse grade. 
74 
Table 4.4. Yield quality of Sellie and PC254K1 inoculated at planting with five geographically-
isolated Ditylenchus africanus populations from Mareetsane (population 1), Jan 
Kempdorp (population 2), Vaalharts (population 3), Schweizer-Renecke 
(population 4) and Theunissen (population 5) in a microplot trial during 2007-
2008. 
Genotype Nematode UBS Choice 
Standard Diverse Crushing 
population (%) (%) (%) (%) (%) 
Sellie 1 14 0 0 61 39 
5 12 0 0 64 36 
2 13 0 0 70 30 
4 12 0 0 58 42 
3 13 0 0 41 59 
PC254K1 4 9 0 41 19 40 
3 4 35 0 21 44 
1 8 0 42 21 37 
2 7 0 33 18 49 
5 9 0 36 19 45 
4.5 DISCUSSION 
No significant interaction between temperature regimes and Pf of the five different D. 
afhcanus populations in the growth cabinet trial implies that the increase in 
population at the different temperature regimes was similar for all. However, the data 
of this trial showed that temperature regime has a great effect on D. africanus 
population growth and that 28 °C is close to the optimum temperature for 
reproduction of this nematode (De Waele & Wilken, 1990) for a variety of different 
populations. The lower Pf at 21 °C and at 35 °C compared to that of 28 °C may be 
attributed to a reduction in egg production and development (De Waele & Wilken, 
1990). 
In the greenhouse trial the respective D. africanus populations behaved differently on 
Sellie than on PC254K1. Although the latter genotype consistently showed resistance 
to all the D. africanus populations its expression of resistance against population 2 
(Jan Kempdorp) was significantly stronger than against the rest of the populations. 
The lack of significant interaction between the two groundnut genotypes and the five 
D. africanus populations implies that the increase of the D. africanus populations on 
each of the two groundnut genotypes were similar. In contrast to the greenhouse 
75 
trial, the five D. africanus populations did not behave differently in terms of 
reproduction rate on Sellie and PC254K1 in the microplqts since no significant 
interaction existed between the two groundnut genotypes and the five populations. 
The lower nematode numbers from Sellie in the microplot trial compared to that 
extracted from Sellie from the greenhouse trial has previously been reported (Venter 
et ai, 1991; Mc Donald et at., 2005). However, the low Pf of the nematodes extracted 
from pods of PC254K1 in the greenhouse and microplot trial confirmed PC254K1's 
resistance to all five D. africanus populations. Resistance of a host plant is 
determined, among others by its effect on nematode reproduction (Trudgill, 1986) 
that could be defined as the ability to inhibit the reproduction of a population relative 
to the reproduction of the population on the susceptible host (Cook & Evans, 1987; 
Roberts, 2002). 
Although the RF of the D. africanus populations on groundnut callus tissue at 28 °C 
in the growth cabinet trial was much higher than those at 21 °C and 35 °C the RF, 
which was also higher than one at the latter two temperature regimes support 
findings of De Waele and Wilken (1990) that egg viability and start of egg hatching at 
these temperatures may be similar to those at 28 °C. This data further indicate that 
all the D. africanus populations were still able to increase at the latter two 
temperatures even though their reproduction rate was not optimal. The resistance of 
PC254K1 to D. africanus was again confirmed by RF in the greenhouse as well as in 
the microplots (Roberts & May, 1986; Windham & Williams, 1988). 
The small range in LIBS % between the five different populations in Sellie indicates 
that the damage potential of the populations on groundnut is similar. Mc Donald et ai 
(2005) showed strong relationships between the presence of D. africanus and UBS 
%. Venter et al. (1991) also provided strong evidence that UBS % is a reliable 
indication of D. africanus damage. The low UBS % of PC254K1 compared to that of 
Sellie is, therefore, further confirmation of its resistance to all five D. africanus 
populations. The differences, though being small, between the UBS % of the 
populations do, however, not agree with differences in Pf or RF of either Sellie 
(susceptible) or PC254K1 (resistant). 
The results of this study indicated that the reproduction and damage potential of the 
five D. africanus populations representative of the local groundnut-production area 
were similar. The near optimum temperature for development of D. africanus 
populations is 28 °C but they are able to reproduce (although not optimally) over a 
76 
wide range of temperature regimes. Resistance of PC254K1 to all five D. africanus 
populations was confirmed in all nematode and plant variables in this study. The 
results suggest that the resistant trait of a D. africanus resistant cultivar developed 
from PC254K1 should be sustainable over the whole groundnut production area. 
77 
CHAPTER 5 
THE MECHANISM OF RESISTANCE TO DITYLENCHUS AFRICANUS 
EXPRESSED BY THE GROUNDNUT GENOTYPE PC254K1 
5.1 INTRODUCTION 
The success of strategies used in long-term nematode management programmes will 
depend on the understanding of the reaction of the host plant to attack by plant-
parasitic nematode populations, including the mechanism of resistance involved 
(Kotcon et a/., 1987). The histopathology of D. africanus has previously been studied 
on the cultivar Sellie, which is highly susceptible to this nematode (Jones & De 
Waele, 1990; Venter et a/., 1995). Resistance to this nematode has been identified 
for the first time during this study (Chapter 3). Therefore, no information is available 
on the histopathology of this nematode on resistant genotypes. The objective of this 
study was to study the histopathology of D. africanus on PC254K1 and compare it to 
that of Sellie in order to determine the mechanism of resistance involved. 
5.2 MATERIALS AND METHODS 
PC254K1 and Sellie material for the histological study was propagated in a 
greenhouse under conditions of optimum growth for the genotypes and those that 
allowed uninhibited development and feeding of D. africanus on its host plant. These 
plants were propagated in the greenhouse from May to October 2008. The 
microscope slides prepared from the PC254K1 and Sellie material were studied 
under a light microscope. 
5.2.1 Trial layout in the greenhouse 
Twenty-eight plastic pots were filled with 4 000 cm3 fumigated, sandy-loam soil as 
described in Chapter 2. Nutrients mixed into the soil (Chapter 2) included 5.80 g 
dolomitic lime, 3.45 g super phosphate (10.5 % P), 2.25 g sodium chloride (KCI) and 
1.60 g calcium nitrate (CaN03) per pot. Seeds were obtained from nematode-free 
mircoplot nurseries (Chapter 2) and treated before planting with fungicide and 
nitrogen-fixing bacteria following the procedures described in Chapter 2. One seed 
was planted per pot to a depth of 5 cm (Chapter 2). 
78 
The trial was done in a greenhouse with a temperature regime of 18 °C to 27 °C and 
a 13-h photoperiod. Six pots each of Sellie and PC254K1 were inoculated at planting 
with 0 (nematode-free), 2 000, 5 000 or 7 000 D. africanus of various life stages. 
Nematodes for inoculation were obtained from an in vitro culture of the Vaalharts 
population (Chapters 2 & 3) of D. africanus according to the procedures described in 
Chapter 2. A plastic marker in each pot indicated the groundnut genotype, nematode 
inoculum level and replicate number. The pots were rotated clockwise every fortnight 
to eliminate the effect of temperature or light gradients in the greenhouse. The plants 
were watered by hand three times a week. 
Pegs not yet developed into mature pods as well as mature pods were collected at 
90 days after plant (DAP), 120 DAP and at harvesting (150 DAP). At each sampling 
time two plants from each genotype and inoculum level were separated from the soil 
(Chapter 2), their pegs and pods removed (Chapter 2), mixed by hand and ten 
randomly-picked pegs and pods were collected from each replicate. Pods collected 
at 150 DAP were photographed to compare external D. africanus damage symptoms 
between the two genotypes before fixing the material in paraformaldehyde. This was 
done only at 150 DAP because this is the period when D. africanus populations peak 
in the pods (Basson et a/., 1990). External damage caused by these nematodes is, 
therefore, supposed to be optimal at 150 DAP. The rest of the plant tissue not used 
for histopathology studies was discarded. The material was washed free of soil under 
a gentle stream of running tap water and fixed in 4 % paraformaldehyde contained in 
plastic 250-ml beakers. The material was stored at room temperature for a maximum 
of three days before preparation for light-microscope studies commenced. 
5.2.2 Preparation of plant tissue for light-microscope studies 
The plant material was prepared at room temperature under a flow cabinet. Pieces of 
peg and pod tissue were cut in approximately 2-mm sections and fixed in Todd's 
fixative (pH 7.5) in 20-ml glass flasks for 2 h (Todd, 1986). After 2 h the Todd's 
fixative was carefully removed with a fine glass pipette and replaced with 0.05-M 
sodium cacodilate buffer in which the tissue samples were impregnated for 15 min. 
This process was repeated three more times. The material was post-fixed in 1-% 
osmium tetroxide for 1 h and then washed in three changes of distilled water for 15 
min each. The material was then dehydrated in an ethanol series of 50 %, 70 %, 90 
% and 100 % for 15 min each. The 100-% ethanol was then carefully poured off and 
replaced with 100-% LR White resin in which the material was allowed to be 
79 
impregnated for 10 min. After that the resin was carefully poured off and replaced 
with fresh resin, leaving it for 1 h. The resin was then again carefully poured off and 
replaced with fresh resin in which the plant material was left overnight at 4 °C in a 
refrigerator. The next day the material was finally embedded in fresh resin contained 
in marked gelatine capsules, which were cured overnight in an oven at 60 °C 
(O'Brien & Mc Culiey, 1981). One-um sections (longitudinal and cross sections) of 
the plant material were cut with glass knives on a rotary microtome, floated on 
microscope slides and air-dried. The material on each microscope slide was stained 
with 0.5-% aqeous toluidine blue (Lillie & Fullmer, 1976) and 0.05-% neofuchsin for 3 
min. The remainder of the stain was rinsed off with tap water and the stained plant 
material allowed to dry for 3 min before it was covered with a cover slide. At each 
sampling interval (90, 120 and 150 DAP) 20 microscope slides consisting of 10 slides 
of peg tissue and 10 slides of pod tissue were prepared and examined for each 
genotype at each D. africanus inoculum level. Light micrographs were taken from 
selected slides. Orientation of the material on the microscope slide and identification 
of the different structures were done with the assistance of trained personnel of the 
School of Environmental Sciences and Development as well as the Electron 
microscopy unit at North-West University, Potchefstroom. 
5.3 RESULTS 
5.3.1 External damage symptoms on mature pods collected at 150 DAP 
At 150 DAP damage symptoms were visible on pods of Sellie inoculated with ca. 2 
000, 5 000 as well as 7 000 D. africanus specimens. Damage symptoms on Sellie 
and PC254K1 pods were most severe at the 7 000-nematodes inoculum level (Fig. 
5.1). Upon inspection of Sellie pods the tissue at the connection of the peg to the pod 
appeared brown and necrotic. The veins that extended longitudinally in the exocarp 
just beneath the surface of the pods had a brown discoloration similar to that of the 
tissue at the peg-pod connection. On some pods of Sellie this dark discoloration 
extended over a large portion of the pod surface. At 150 DAP pods of PC254K1 at an 
inoculum level of 7 000 D. africanus specimens were not discolored at the peg-pod 
connection point or over the pod surface (Fig. 5.1). No discoloration of the peg-pod 
connection or the pod surface was visible on Sellie or PC254K1 from the nematode-
free control at 150 DAP. 
80 
Figure 5.1. External symptoms on Setlie and PC254K1 pods at an inoculum level of 
7 000 Ditylenchus africanus specimens 150 DAP in a greenhouse trial 
during 2008. 
5.3.2 Histopathology 
5.3.2.1 Pegs 
The histopathology in longitudinally-sectioned pegs of Sellie at the various inoculum 
levels and different sampling times were similar in appearance to that of 
longitudinally-sectioned pegs from PC254K1 collected at similar inoculum levels and 
sampling times (Fig. 5.2 A & B). Each peg had a distinct exocarp with developing 
vascular bundles in the mesocarp. The inner layer of the mesocarp was 
differentiating into a fibrous mesocarp layer. The cells of the fibres were still thin-
walled and unlignified (Fig. 5.2 A & B). In the central pith of the PC254K1 pegs a part 
of the fruit locule contained the ovule (Fig. 5.2 B). Within the ovule resided the 
embryo sac in which a groundnut embryo would be formed. No nematodes were 
observed in the peg tissue of either Sellte or PC254K1 from the nematode-free 
controls or any of the nematode \noculum \eve\s at 90, 120 or 150 DAP. 
81 
Figure 5.2. Light micrograph of a longitudinal section of a peg from A. Sellie and 
B. PC254K1 at an inoculum level of 7 000 Ditylenchus africanus 
specimens at 150 DAP from a histopathology study during 2008. 
A & B magnified 20 x. D = developing fibre layer, EN = endocarp, 
ES = embryo sac, EX = exocarp, F = fruit locule, M = mesocarp, 
O = ovule, V = vascular bundle. 
82 
Figure 5.3. Light micrograph of a cross section through the vascular region of pegs from 
A. Seilie and B. PC254K1 at an inoculum level of 7 000 Ditylenchus africanus 
specimens at 150 DAP from a histopathology study during 2008. A & B 
magnified 100 x. DUM = damaged undifferentiated tracheary elements of the 
metaxyiem, DP = differentiated tracheary elements of the protoxylem, EN = 
endocarp, M = mesocarp, UM = undifferentiated tracheary elements of the 
83 
metaxylem. 
Cross sections of the vascular region of pegs of inoculated as well as uninoculated 
Sellie and PC254K1 at 90, 120 and 150 DAP were similar in appearance (Fig. 5.3 A 
& B). The vascular regions are situated in the parenchyma of the mesocarp adjacent 
to that of the endocarp. Each vascular region consists of a row of differentiated 
tracheary elements of the protoxylem that is thick-walled and has a ribbed 
appearance. Adjacent to the differentiated tracheary elements of the protoxylem the 
undifferentiated tracheary elements of the metaxylem consist of a long row of thin-
walled cells. A number of cells along the undifferentiated tracheary elements of the 
metaxylem appear damaged and the cell walls between these cells are broken down. 
This damage along the undifferentiated tracheary elements of the metaxylem was 
also visible in Seiiie and PC254K1 plants from the nematode-free contro\ (not 
shown). 
5.3.2.2 Pods 
From 90 DAP D. africanus was observed in pod tissue of Sellie at all three different 
inoculum levels. At 150 DAP several nematodes were observed in Sellie pods at the 
2 000, 5 000 and 7 000 inoculum levels. No nematodes were, however, observed in 
pods of Sellie from the nematode-free control at 90 and 150 DAP. Cross sections of 
mature pods of Sellie 150 DAP inoculated with ca 7 000 nematodes are presented in 
Figure 5.4 A & B. The funicle is surrounded by parenchyma cells of the endocarp 
(Fig. 5.4 A). Adjacent to the endocarp the cells of the fibrous layer of the mesocarp 
are thick-walled. Parenchyma cells of the mesocarp are located between the fibrous 
mesocarp and the corky )ayer of the exocarp. Nematodes were not present in the 
thick-walled cells of the \/ascv\ar bundle of the funicle, in the thick walled cells of the 
fibrous mesocarp or in the cells of the exocarp. They were observed in the 
parenchyma cells of the mesocarp as well as in those of the endocarp (Fig. 5.4 A). 
The largest proportion of the nematodes present in Sellie pods was observed around 
the vascular bundle of the funicle and in the parenchyma cells of the endocarp (Fig. 
5.4 B). Walls of the malformed, elongated parenchyma cells around the nematodes 
that are located in the endocarp around the vascular bundle are broken down and 
cells started to aggregate. In advanced stages of parenchyma damage the cells 
completely collapse (Fig. 5.4 B). These cells were not malformed, aggregated or 
collapsed in pod tissue of uninoculated Sellie plants from the nematode-free control. 
84 
1/ w -
? - W£w' 
N . . ? '%/ 
S3 . RY«N Q ? 
000 pm 
3 
; 
>rJ -
V. 
, 
Figure 5.4. Light micrograph of a cross section through A. The vascular bundle of 
the funicle and surrounding endocarp and mesocarp and B. The vascular 
bundle of the funicle and surrounding endocarp of a mature, Sellte pod at 
an inoculum level of 7 000 Ditylenchus africanus specimens at 150 DAP 
from a histopathoiogical study during 2008. A magnified 20 x and B 40 x. 
EN = endocarp, EX = exocarp, FM = fibrous mesocarp, M = mesocarp, 
85 
N = nematocie, V - vascular bundle of the funicle. 
Nematodes were observed in pods of PC254K1 for the 2 000, 5 000 and 7 000 
inoculum levels at 90 DAP. No nematodes occurred in pods of PC254K1 and Sellie 
of the nematode-free control plants. Cross sections of the vascular region and 
surrounding mesocarp of a mature PC254K1 pod collected at 150 DAP from plants 
inoculated with ca 7 000 nematodes are presented in Figure 5.5 A & B. The vascular 
bundles are located in the mesocarp adjacent to the fibrous mesocarp, which 
consists of thick-walled cells. In contrast to the high numbers of nematodes observed 
in pod tissue of Sellie, few nematodes were seen in the pod tissue of PC254K1 (Fig. 
5.5 A & B). D. afhcanus was not observed in the parenchyma cells of the endocarp, 
the thick-walled cells of the vascular bundles or in the thick-walled cells of the fibrous 
mesocarp of the latter genotype. They were observed only in cells in the mesocarp 
immediately adjacent to the thick-walled vascular bundles (Fig. 5.5 B). 
86 
Figure 5.5. Light micrograph of a cross section through A. The vascular region and 
surrounding mesocarp and B. The vascular region of a mature 
PC254K1 pod at an inoculum level of 7 000 Ditylenchus africanus 
specimens at 150 DAP from a histopathology study during 2008. 
A magnified 20 x and B 40 x. EN = endocarp, FM = fibrous mesocarp: 
M = mesocarp, N = nematode, V = vascular bundles. 
87 
A region around the vascular bundles of a mature pod of PC254K1 at 150 DAP 
inoculated with ca 7 000 D. africanus specimens is presented in Fig. 5.6. 
Figure 5.6. Light micrograph of a cross section through the cells of the vascular 
region of a mature PC254K1 pod at an inoculum level of 7 000 
Ditylenchus africanus specimens at 150 DAP from a histopathology 
study during 2008. Magnified 100 x. DUM = damaged undifferentiated 
tracheary elements of the metaxylem, DP = differentiated tracheary 
elements of the protoxylem, M = mesocarp, N = nematode. 
Differentiated tracheary elements of the protoxylem are thick-walled with a ribbed 
appearance similar to those in the peg tissue (Fig. 5.3 A & B). D. africanus was 
observed only in the area of the undifferentiated tracheary elements of the 
protoxylem. Parenchyma cells around the undifferentiated tracheary elements of the 
protoxylem were nematode-free, not malformed, aggregated or coWapsed such as 
that of inoculated Sellie. 
88 
5.4 DISCUSSION 
The brown discoloration on the pod surface and the necrotic tissue at the connection 
point of the peg to the pod observed on D. africanusAnfected pods of Sellie during 
this study was similar to those observed on the same cultivar in previous studies 
(Jones & De Waele, 1988; De Waele et ai, 1989). Contrary to the situation with pods 
of Sellie, those of inoculated PC254K1 showed no external damage symptoms 
associated with D. africanus, confirming the resistance of this groundnut genotype to 
D. africanus even at high inoculum levels (also see Chapter 3). 
Absence of D. africanus from peg tissue of inoculated Sellie of this study is in 
agreement with previous studies (Venter et a/., 1995). It is unclear what caused the 
damage to undifferentiated tracheary elements of the metaxylem in the pegs of 
inoculated Sellie as well as PC254K1 because similar damage was present in the 
same region in peg tissue of their uninoculated counterparts. 
The damage caused by D. africanus in infected, mature Sellie pods at cellular level 
as well as the fact that individuals were present between the parenchyma cells of the 
mesocarp and endocarp but not beyond the mesocarp support observations made in 
previous studies (Jones & De Waele, 1988; De Waele et ai, 1989). Jones and De 
Waele (1990) and Venter et al. (1995) suggested that broken-down cell walls as 
observed in our study is a result of enzyme secretions of D. africanus. Seinhorst 
(1957) proposed that for proper development of D. dipsaci in host tissues, plant cells 
must be destroyed by dissolution of the middle lamella through enzymatic secretions 
by nematodes. In our study the macerated cells observed throughout the meso- and 
endocarp of infected Sellie pods indicated that this may also apply to D. africanus. 
Lack of damage and the absence of D. africanus in pod tissue of PC254K1 confirmed 
the genotype's resistance and suggests that the mechanism of resistance may be 
inhibition of proper development and subsequent reproduction of D. africanus. The 
few nematodes found in the region of the undifferentiated tracheary elements of the 
metaxylem seemed not able to induce dissolving of the cell walls and the middle 
lamella of the neighbouring parenchyma cells (Seinhorst, 1957). As trachery 
elements mature, they lose their end walls through hydrolysis of the unlignified 
intercellular layer and their nuclei and cell contents, leaving hollow, dead cells that 
form vessels or tracheids (Fukuda, 1997). It appears as if hydrolysis of the walls by 
PC254K1 plants enabled a small number of D. africanus individuals to survive in 
89 
these tracheary elements. Therefore, although PC254K1 is not immune to D. 
africanus this genotype has the ability to prevent optimum development and 
reproduction of this nematode in its pods. This reaction of PC254K1 to D. africanus 
development and reproduction implies that antibiosis is most probably the 
mechanism of resistance involved. 
90 
CHAPTER 6 
ORIGIN OF RESISTANCE TO DITYLENCHUS AFRICANUS IN PC254K1 AND 
IDENTIFICATION OF MOLECULAR MARKERS 
6.1 INTRODUCTION 
The groundnut breeding line PC254K1 has been identified during this study as a 
source with a high level of resistance to D. africanus and may have potential to be 
used in groundnut-breeding programmes (Chapters 3 to 5). Once a source of 
resistance has been identified a plant breeder needs to know the manner in which 
the trait is inherited (monogenic, oligogenic or polygenic) (Roberts, 1990) so that the 
most efficient breeding strategies could be selected for providing a desirable product 
that could enter the seed market by the shortest possible route (Boerma & Hussey, 
1992). 
Molecular markers could play an important role when improvement of the efficiency 
and duration of conventional plant breeding programmes is desired (Ortiz, 1998; 
Kasha, 1999). Markers that are closely linked to specific genes could facilitate 
verification of the presence of a specific trait as well as pyramiding of a number of 
desired genes into a single variety (Mienie et al., 2005). The use of molecular 
markers is, therefore, widely accepted as valuable tools for the genetic improvement 
of various crops (Collard et al., 2005). Church et al. (2001) identified the RFLP 
markers R2430E and S1018E that flanked a dominant resistance gene locus for root-
knot nematode resistance in groundnut but since the resistance of PC254K1 to the 
groundnut pod nematode has only been identified during this study (Chapter 3), no 
information is available on markers associated with this specific trait. 
Therefore, the objectives of this part of the study were to i) establish the mechanism 
of inheritance of the D. africanus resistance trait in PC254K1 and to ii) identify 
molecular markers linked to this trait. 
91 
6.2 MATERIALS AND METHODS 
6.2.1 Origin of the resistant trait 
6.2.1.1 First-generation crosses (Fi seed) 
This study was conducted in a greenhouse with a 18 °C to 30 °C temperature regime 
and a 13-h photoperiod. A total of eighteen 25 000-cm3 plastic pots were filled with 
fumigated, sandy-loam soil (Chapter 2). The following nutrients were added to each 
pot: 36.5 g dolomitic lime, 22 g super phosphate (10.5 %), 14.5 g sodium chloride 
(KCI) and 9.4 g calcium nitrate (CaN03) mixed into the soil as described in Chapter 
2. 
Sellie and PC254K1 seed were obtained from nematode-free mircoplot nurseries and 
treated before planting with fungicide and nitrogen-fixing bacteria (Chapter 2). Plants 
used as male parents in the crosses were planted in two 1-week-consecutive 
batches for continuous availability of pollen for artificial pollination of female plants 
throughout the flowering phase. Each batch of males consisted of three pots planted 
with Sellie and three pots of PC254K1. The first batch was planted a week before 
planting of the females. A week later three pots each of Sellie and PC254K1 were 
planted to be used in the crosses as females and three pots each of Sellie and 
PC254K1 were planted to be used in the crosses as males. Female plants were kept 
apart from male plants. A plastic marker placed in each pot indicated the relevant 
groundnut genotype and whether it was used as female or male parent. 
Crosses with Sellie as the male and PC254K1 as the female and vice versa 
(reciprocal crosses) were made as soon as the 18 female plants (nine Sellie and nine 
PC254K1 plants) started flowering approximately five weeks after planting. Young 
flowers situated on the first three nodes of the first three branches of female plants 
were used for the crosses because these parts carry approximately 74 % of the total 
number of pods at harvesting, which in turn produces approximately 92 % of the total 
kernel yield of a groundnut plant (Swanevelder, 1997). Flowers in these regions that 
were not used in crosses as well as those that developed elsewhere on the plants 
were removed on a daily basis to prevent undesirable self-pollinations. Crosses were 
made daily on flowers that have developed the previous day. Self-pollination of the 
developing flowers before artificial pollination was avoided by removing the anthers 
between 15:00 and 18:00 the day before maturation of each flower's own stigmata 
92 
(Knauft & Wynne, 1995). The lower lip of the calyx, wing and keel petals were 
removed with a sharply-pointed forceps to expose the anthers and stigmata. The 
anthers were then carefully removed without damaging the stigmata. Emasculated 
flowers were marked with pieces of coloured string, loosely tied around them so that 
they could be identified for artificial pollination the following day (Fig. 6.1). 
Figure 6.1. Emasculated flowers on a groundnut plant marked 
with coloured strings for artificial pollination. 
Artificial pollination of flowers proceeded between 7:00 and 9:00 in the morning 
following emasculation because this is the period during the day when the stigmatic 
tissue is most receptive and pollen is most viable (Knauft & Wynne, 1995). Po\)en 
was harvested from anther sacs of mature flowers on male parents just before 
pollination. A flower was removed from a male parent and ripe pollen was carefully 
squeezed from the anther sacs onto a forceps. The pollen was then carefully 
transferred onto a receptive stigma of an emasculated flower marked with a coloured 
string. The forceps used during artificial pollination was dipped in 70 % ethanol after 
each pollen transfer in order to prevent cross-pollination. To prevent dehydration of 
stigmata that were exposed during emasculation and of the pollen being transferred 
to the stigmata of female parents the latter flowers were covered with plastic bags 
after pollination. The plastic bags were each supported by 60 X 36-cm2 wire frames 
(Fig. 6.2). 
93 
Figure 6.2. Female plants covered with a plastic bag supported 
by a wire frame. 
Pegs that developed from 40 successfully pollinated flowers seven to 10 days after 
pollination were labelled to indicate the female and male parent. Mature pods that 
developed approximately five months after artificial pollination were removed from 
plants, placed in appropriately marked brown paper bags and were air-dried in a 
greenhouse for seven days. After this the pods were shelled to collect Fi seeds. 
Twenty Fi seeds were obtained from Seliie (<$) and PC254K1 (2) and its reciprocal 
crosses, respectively. 
6.2.1.2 Second-generation (F2) seed 
Seed of fi from the reciprocal Seliie and PC254K1 crosses was planted in a 
greenhouse specially adapted for breeding purposes (Fig. 6.3) during 2005-2006. 
The four 2.1 x 1.4 x 1.4-m3 brick troughs are built over a drainage system to prevent 
waterlogging. Each trough was filled with EDB-fumigated, sandy-ioam soil that was 
re-fumigated before planting of the trial (Chapter 2). Nutrient amendments consisting 
of 200 g dolomitic lime, 139 g super phosphate (10.5 % P), 42 g KCI and 40 g CaN03 
per trough were incorporated into the top 30 cm of the soil of each of the troughs 
following procedures described in Chapter 2. 
94 
Figure 6.3. Groundnut breeding facility in a greenhouse. 
Seed was treated with fungicide and inoculated with nitrogen-fixing bacteria (Chapter 
2) before planting at an intra-row spacing of 10 cm and inter-row spacing of 45 cm 
(Chapter 2). In the first four adjacent rows the 20 F, seed from Sellie ($) x PC254K1 
(3) crosses were planted and in the last four rows the 20 F, seed from PC254K1 ($) 
x Sellie (<$) crosses. Plastic markers placed in front of the first row of each four-row 
block indicated the female and male parent. Plants were allowed to self-pollinate and 
were watered three times a week by hand. At harvesting the plants were lifted from 
the soil (Chapter 2) and the pods (Fig. 6.4) removed. 
Figure 6.4. Mature pods produced by FT plants after crosses 
were made between Seliie and PC254K1. 
95 
The pods were placed in marked paper bags and air-dried in a greenhouse for seven 
days. After this they where shelled to obtain F2 seed. Hundred and sixty-three seeds 
were obtained from the F1 plants from Sellie {$) x PC254K1 (-) crosses and 159 
seeds form the F, plants of PC254K1 (?) x Sellie (c?). 
6.2.1.3 Evaluation of the F2 progeny for D. africanus resistance 
During 2006-2007 the abovementioned F2 progenies were evaluated for resistance to 
D, africanus in a microplot facility that consists of a row of 20 rectangular ciay-brick 
troughs (Chapter 2). The troughs were filled with the same EDB fumigated, sandy-
loam soil used in the pots in the greenhouse in this chapter (Chapter 2). Nutrient 
amendments per plot were 75 g dolomitic lime, 136 g super phosphate (10.5 % P), 
40 g KCI and 41 g CaN03. The nutrients were incorporated into the top 30 cm of the 
soil as described in Chapter 2. 
The trial was planted in a randomised complete block design. The four treatments 
consisted of the parents Sellie (susceptible) and PC254K1 (resistant), F2 from Sellie 
($) x PC254K1 (oA) cross and F2 from PC254K1 (?) x Sellie (<J) cross. Four rows 
were planted in each trough at an intra-row spacing of 5 cm and an inter-row spacing 
of 45 cm (Chapter 2). Each row represented a treatment and contained 20 seeds. 
The F2 seed was treated with fungicide and inoculated with B. arachis (Chapter 2). 
White plastic markers at the front end of each row indicated the treatment. 
At planting each seed was inoculated with + 3 000 D. africanus of various life stages. 
The nematodes for inoculation were extracted from D. africanus cultures and 
prepared for inoculation as described in Chapter 2. The trial was irrigated 
supplementary to rainfall (Chapter 2). Each plant was labelled after germination (see 
also 6.2.2.1). 
Nematode assessments were done at harvesting on all F2 plants. Nematode 
extractions were done from peg-, hull- and kernel-samples from each plant, counted 
under a research microscope and expressed as number of Pf per 15 g pods (Chapter 
2). 
96 
6.2.2 Molecular marker identification 
6.2.2.1 DNA extraction 
Three weeks after germination the F2 seedlings (6.2.1.3) from the PC254K1 {$) x 
Sellie (<$) cross were each labelled and a unique number was allocated to each 
seedling. Eight leaves were sampled per plant and placed in appropriately marked 2-
ml Eppendorf tubes. They were freeze-dried and stored at -20 °C until extraction of 
their DNA. 
Total genomic DNA was isolated from the leaves using a modified 
hexadecyitrimethylarnmoniurn bromide (CTAB)-method (Saghai-Maroof ef al., 1984). 
Freeze-dried leaves of each plant were ground to a fine powder and sub-sampled to 
approximately 250 pi of fine leaf powder. Each sub-sample was then transferred to a 
2-ml microfuge tube, topped with 700 pi CTAB buffer consisting of 100 mM 
Tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCI) (pH 8.0), 20 mM 
ethylenediaminetetra acetic acid (EDTA) (pH 8.0), 1.4 M sodium chloride (NaCI), 2 % 
(w/v) CTAB and 0.2 % (v/v) p-mercaptho-ethanol and incubated at 65 °C for 1 h. 
After incubation the suspension (aqueous phase) was extracted with a 500 ul 
chloroforrrvisoamylalcohol solution (24:1 v/v). The aqueous and chloroform phases 
were separated by centrifugation of the suspension at 12 000 r.p.m. for 10 min. The 
chloroform phase was discarded and 0.02 Mg.pL"1 ribonuclease A (RNase A) was 
added to the aqueous phase contained in a microfuge tube and then incubated for 30 
min at room temperature to digest RNA that could contaminate the DNA. DNA was 
centrifuged at 12 000 r.p.m. for 15 min and precipitated from the aqueous phase with 
Yz volume isopropanol. The precipitate collected in the microfuge tube was washed 
with 500 pi ice-cold, 70 % (v/v) ethanol and then centrifuged at 12 000 r.p.m. for 5 
min. The pellet was air-dried for one hour and then resuspended in Tris-EDTA (TE) 
buffer (10 mM Tris-HCI, pH 8.0; 1 mM EDTA, pH 8.0). Absorbencies were measured 
at 260 nm and 280 nm to determine both DNA quantity and quality. 
6.2.2.2 Amplified fragment length polymorphism (AFLP) analysis 
AFLP analysis was conducted using a protocol modified after Vos ef al. (1995). The 
primer pair combinations used to perform the analyses is presented in Table 6.1. 
Mlu\ primers (Ml) were screened in combination with Msel-primers (M). The codes 
following Ml or M refer to the three selective nucleotides at the 3'-end of the primer. 
97 
Tabie 6.1. Mlu\ (Ml) and Mse\ (M) adaptor, primer+1 and primer+3 sequences used in 
amplified fragment length polymorphism (AFLP) analyses of genomic 
deoxyribonucleic acid (DNA) of PC254K1, Sellie and the F2 generation resulting 
from a PC254K1 x Sellie cross. 
Enzyme Type Sequence (5* - 3') 
~Ml Adaptor-F CTCGTAGACTGCGTAAC 
Adaptor-
CGCGGTTACGCAGTC 
R 
M Adaptor-F GACGATGAGTCCTGAG 
Adaptor-
TACTCAGGACTCAT 
R 
Ml Primer+1 GACTGCGTAACCGCGT 
Primer+3 GACTGCGTAACCGCGTNN 
TNN = TTA, TCA, TGC, TTA, TCT, TGT, TTG 
M Primer+1 GATGAGTCCTGAGTAAN 
N = A, C, G, T 
Primer+3 GATGAGTCCTGAGTAAN NN 
NNN = ACA, ACC, ACT, CAA, CAC, CAG, CAT, CCG, CGT, CTA, 
CTC, CTG, 
CTT, GAA, GAG, GGC, TAC, TTG 
Primers and adaptors were synthesised by Applied Biosystems (Integrated DNA 
Techologies, UK). Ml primers were fluorescently labelled using 6-carboxyfluorescein 
(6-FAM), NED™, VIC™ or PET™. Adapters were prepared as follows: Equimolar 
amounts of each relevant pair (Table 6.1) were combined in a 0.2-ml Eppendorf tube. 
The solution was heated for 10 min to 65 °C in a water bath and then allowed to cool 
down to room temperature to ligate the primers. 
Genomic DNA (500 ng) was digested for 5 h at 37 °C using four units (U) Mse\ in 
NEB Buffer2 (50 mM NaCI, 10 mM Tris-HCl, 10 mM magnesium chloride (MgCI2), 1 
mM dithiotreitol (DTT), pH 7.9; New England Bioiabs), followed by the addition of 50 
mM NaCI and 5 U Miu\ before overnight incubation at 37 °C. Adapters were ligated to 
the restricted genomic DNA by addition of 50 pmol Mse\ adapter, 5 pmol of Mlu\ 
adapter and 1 U T4 DNA ligase. Ligation was allowed overnight at 16 °C. 
Pre-ampiification reactions were performed in 50-ul reaction mixtures consisting of 5 
ul ligated DNA, 200 uM of each dNTP, 30 ng of each pre-ampliftcation primer (Ml-
and M-phmer+1) (Table 6.1), 1x Taq polymerase buffer, 2 mM MgCI2and 1 U Taq 
98 
DNA polymerase. Samples were amplified in a Thermo Electron Multiblock System 
using 30 cycles of 30 s at 94 °C, 1 minute at 56 °C and 1 min at 72 °C each. Quality 
and quantity of pre-amplification products were determined by electrophoresis in a 
1.5 % (m/v) agarose gel and diluted 1:50 prior to selective amplification. 
Selective amplification reactions were performed in a 20-ul reaction volume 
consisting of 5 uL of 50-fold diluted pre-amplification product, 30 ng MI+3 primer 
(Table 6.1), 30 ng M+3 primer, 2 mM MgCI2l 100 ug.mL"1 bovine serum albumin 
(BSA), Taq polymerase buffer and 0.02 U.uL"1 Taq DNA polymerase. Samples were 
amplified for one cycle of 30 s at 94 °C, 30 s at 65 °C and 1 min at 72 °C. The 
annealing temperature was lowered 1 °C for each of nine cycles followed by 24 
cycles of 30 s at 94 °C, 30 s at 56 °C and 1 min at 72 °C each. 
Capillary electrophoresis was done on an ABI3130x DNA sequencer with 16 
capillaries. Diluted (1:20) polymerase chain reaction (PCR) amplification products 
generated by three different primer combinations were mixed and loaded together 
with a GeneScan™-500 LIZ® size standard in HiDi formamide onto the ABI3130x/. 
DNA fragments resulting from the primer combinations were distinguished using the 
distinct emission spectra of the fluorescently-labelled Ml primers, while the samples 
were separated in the same electrophoresis capillary. Analysis was performed using 
the GeneMapper Version 4.0 programme. 
6.2.2.3 Bulk segregant analysis 
This part of the study was done after nematode evaluations (6.2.1.3) because the 
percentage reproduction rate (RR) of the nematodes was used to distinguish 
between resistant (RR was less than 10 % of that of Sellie), segregating (RR was 
between 10 % and 100 % of that on Sellie) and susceptible (RR was 100 % or more 
than that of Sellie) F2 plants (also see 6.3.1) to support this analysis. A total of 107 
primer combinations were tested on the DNA from leaves of the individual plants of 
the two parents and bulks. The two bulks consisted of DNA extracted from the leaves 
collected from six F2 plants that showed the highest resistance levels to D. africanus 
(RR < 10 % of that on Sellie) and that of leaves collected from six F2 plants that 
showed susceptibility to D. africanus (RR > 100 % of that on Sellie). Primer pair 
combinations that generated informative polymorphisms between the parents and 
bulks (fragments present in resistant parent and bulk or in susceptible parent and 
99 
bulk) were tested on ali of the individual F2 plants (103) from the segregating 
population (RR were between 10 % and 100 % of that in Sellie). 
PCR reactions on the 103 individual plants were done separately and then 
multiplexed by combining 1 pi of each of three different-coloured dyes for the same 
individual in 20 pi of 0.1 xTE buffer. Diluted PCR multiplex reactions (1 pi) were 
added to 9 pE HiDi-formide/GS500LIZ contained in a 96-well microplate, denatured at 
95 °C for 3 min and immediately cooled down by placing the microplate on ice. The 
plate was covered with an ABI3130 septum and briefly centrifuged at 12 000 r.p.m. 
before loaded onto the ABI3130x/for electrophoresis. 
6.3 STATISTICAL ANALYSES 
6.3.1 Origin of the resistant trait 
RR of D. africanus was determined on each F2 plant in the microplot trial to express 
the level of resistance (Timper et al., 2003) and to separate resistant F2 plants from 
susceptible ones (Church et ai., 2005). The equation used was RR = Pf (F2) / Pf 
(Sellie) x 100. For preliminary hereditary evaluations the RR of the nematodes on the 
F2 plants was combined in resistant (RR < 10 % of that on Sellie) (Abdel-Momen et 
ai., 1998; Hussey & Janssen, 2002; Timper et ai, 2003), segregating (RR was 
between 10 % and 100 % than that on Sellie) and susceptible (RR > 100 % than that 
of Sellie) to support bulk-segregant analyses (6.2.2.3). An incidence plot (Stat 
Graphics 5 Plus for Windows) was used to determine the distribution in the F2 
mapping population in terms of Pf in pods. 
6.3.2 Marker identification 
A binary matrix that reflected specific AFLP markers as present (1) or absent (0) was 
generated for each genotype using GeneMapper 4.0. Only markers that were reliable 
(association between the marker and D. africanus resistance trait was acceptable if 
the probability was P < 0.05) and repeatable were considered. The association 
between a marker and the D. africanus resistance trait was determined by doing a 
simple regression analysis for the F2-mapping generation and dominant loci (genetic 
markers that occupy specific genomic positions within chromosomes) (AFLP 
analysis). Regression analyses of the data from the AFLP analysis were done using 
a linear model from StatGraphics Plus 5 for Windows. The genetic marker data 
100 
served as the independent variable and D. africanus numbers per 15 g pod (Pf) as 
the dependent variable. Association between the marker and D. africanus resistance 
trait was considered significant if the probability was P < 0.05. The R-squared 
statistic (R2) was used as a measure of the magnitude of the association. 
A Chi-square (Chi2) analysis was performed to test distortion in segregation of the 
markers. Linkage analysis and drawing of the linkage map was done using 
MAPMAKER.EXP 3.0 (Lander ef a/., 1987). MAPMAKER.QTL 1.1 (Paterson ef a/., 
1988) was used for the localisation of the resistance gene/quantitative trait loci (QTL) 
on the linkage map in a recessive model (Lincoln ef a/., 1992) to scan linkage groups 
(LG) for the presence of QTL effects at a log-likelihood score (LOD) threshold of 3.0 
in every 2.0 centi Morgan (cM) interval. A LG indicated the position and relative 
genetic distances in cM between the markers along the particular chromosome 
involved. The most important use of the constructed linkage map was to identify 
chromosomal locations of genes and QTLs (Collard ef a/., 2005) associated with 
resistance to D. africanus and to indicate the position and relative genetic distances 
between markers along chromosomes (Collard ef a/., 2005). Genes that are closely 
together or tightly linked will be transmitted together more frequently from parent to 
progeny than genes or markers that are located further apart (Collard ef a/., 2005). 
The Haldane mapping function was used to assign markers to linkage groups. The 
position of the resistance gene on the map was determined by treating the trait as a 
marker and mapping it with other markers or by identifying the position of the 
resistance gene by using the scan command of MAPMAKER.QTL 1.1 as described 
by Herselman et al. (2004). Markers were assigned to linkage groups if the LOD < 
3.0 and maximum recombination frequencies (9) were < 0.5. 
6.4 RESULTS 
6.4.1 Origin of the resistant trait 
A total of 163 F2 seeds resulted from the Sellie ($) x PC254K1 (o) cross in the 
greenhouse. D. africanus numbers (Pf) in the pods of the F2 plants ranged from zero 
to 6 500 nematodes per plant. Forty-three F2 plants showed resistance (RR < 10 %) 
and 37 F2were susceptible (RR > 100 %) (Table 6.2). The RR of 83 F2 plants ranged 
between 11 and 99 % and could be classified as an intermediate group 
(segregating). 
tOl 
The F2-plant population from the PC254K1 (9) X Sellie (c?) cross consisted of 159 
plants. Pf on the F2 plants from the latter cross ranged from zero to 7 756 per plant. 
Fifty-one F2 plants showed resistance (RR £ 10 %) and 35 F2 plants were susceptible 
(RR > 100 %) (Table 6.2). RR of the segregating F2 plants (73) ranged between 11 % 
and 99 % and could be classified as an intermediate group. 
Table 6.2. Percentage reproduction rate (RR) of Ditylenchus africanus on pods of F2 plants 
from Sellie (I) x PC254K1 (o) cross and its reciprocal PC254K1(9) x Sellie (<5) 
from a microplot trial during 2006-2007. 
, Sellie' (:?} x PC254K1J (<?) PC254K1(Q) x Sellie (d1) RR 
No of F2 plants No of F2 plants 
51 
33 
12 
9 
1 
3 
5 
2 
35 
< 10 43 
<20 27 
<30 24 
<40 8 
< 50 9 
<60 5 
<70 4 
<80 3 
<90 3 
> 100 37 
1 RR = Pf(F
2)/Pf (Sellie) x 100 
2 Susceptible parent 
3 Resistant parent 
When the F2 plants were subjected to an incidence plot the distribution did not match 
the Mendelian laws for heredity (Klug & Cummings, 1994) for the Sellie (9) x 
PC254K1 (<3) cross (Fig. 6.5) or its reciprocal (Figs 6.5 & 6.6). These plots showed a 
normal distribution in the F2 plant population in terms of Pf. 
102 
25 
] 
0 
E 
Pf/ l5gpod[ln(x-1)] 
Figure 6,5. Distribution of F2 plants from a cross between susceptible Sellie (2) and 
resistant PC254K1 (c?) according to the mean number of Ditylenchus 
africanus ln(x+1) {Pf) in pods per plant at harvesting of a microplot trial 
during 2006-2007. 
30 
25 
20 
o 15 
! 
10 
? 
I JHHIMIHI 
Pf M5gpod(ln(x*1)] 
Figure 6,6. Distribution of F2 plants from a cross between resistant PC254K1 {$) and 
susceptible Sellie ($) according to the mean number of Ditylenchus 
africanus ln(x+1) (Pf) in pods per plant at harvesting of a microplot trial 
during 2006-2007. 
103 
6.4.2 Molecular marker identification 
6.4.2.1 AFLP analysis 
Results obtained from the AFLP primers tested on the DNA of the parents and F2 
bulks are provided in Table 6.3. 
Table 6.3. AFLP primers tested on the DNA of the parents PC254K1 (resistant) (?) and 
Sellie (susceptible) (<$) and the F2 progeny from PC254K1 x Sellie. 
Mlu\-Mse\ I reactions 
Primer combinations tested 107 
Total polymorphisms 430 
Informative primers between parents and bulks 23 
Average polymorphisms / primer combination 20.5 
Maximum polymorphisms / primer combination 37 
Minimum polymorphisms / primer combination 4 
Hundred-and-seven primer combinations were tested during this study and a total of 
430 polymorphisms were detected (Table 6.3). The polymorphisms included a 
minimum of four and a maximum of 37 polymorphisms per primer combination. 
Twenty-three of the 107 AFLP primers tested on the DNA of the parents and F2 bulks 
were informative. These AFLP primers corresponded to fragments in the DNA from 
the resistant parent and resistant bulk of F2 plants or to fragments in the DNA from 
the susceptible parent and susceptible bulk of F2 plants. These informative AFLP 
primers were then further tested on the DNA of the 12 individual F2 plants from the 
two bulks (6.2.2.3). Nine of the 23 informative AFLP primer combinations showed to 
be most useful and were tested on the entire population of 103 individual plants 
consisting of the 12 plants from the two bulks and 91 plants from the rest of the 
segregating population. 
6.4.2.2 Linkage analysis 
According to the Chi2 test performed on the data 84 loci segregated into the expected 
3 : 1 pattern (P < 0.05) (data not shown). These 84 loci were used for linkage 
mapping (Lander et al., 1987). The nine primer combinations detected 
polymorphisms in the 91 plants of the segregating F2from the PC254K1 ($) x Sellie 
104 
(S) cross, which were used for the construction of a putative linkage map. Seven 
LGs were mapped (Fig. 6.7). 
LG1 LG2 LG3 LG4 
o.o-
45.2 
67.4-
76.7-
100.1 
144.0 
178.7 
223.7 
TTACTC101 
TCAGAA103 
TCAGAA89 
TAAGGC114 
- TCTCGT94 
TCTGAA83 
0.0 
36.5 
44.2 
53.9 
69.2 
80.6 
Pr TCTCTA154 
TAAGGC79 
TAAGGC66 
TAAGGC69 
- TAAGGC27 
- TAAGGC15 
0.0 
32.1 
47.7-
67.5-
87.0 
105.7 -
140.2 
TCAACT35 o.o-f ¥- TCAACT442 
15.5 -t \~ TCTGAG414 
TCTGAG354 
TCTGAG203 
TCTGAG218 
TCTGAG206 
TCTGAG197 
TCTGAG490 
LG 
5 
LG 
6 
LG7 
0. 
0 
33.2 
64. 
8 
TCAGAA19 
0 
TCAGAA23 
2 
TCAGAA35 
5 
0. 
0 
25.5 
72.8 
118.2 
148.7-
206. 
4 
TCTCAT13 
6 
TCTCAT13 
7 
TCTCAT10 
3 
TCTCGT15 
0 
TCAGAA63 
TCTCTA119 
0.0 
31.7 
70.8 
91.1 H 
99.2 
113.0 
141.3 
197.3 
TGTCCG141 
TGTCCG138 
— TGTCCG198 
4- TGTCCG84 
TGTCCG88 
-h TGTCCG68 
TCTCTA163 
TGTCCG98 
Figure 6.7. AFLP-based, genetic-linkage map developed for groundnut using a F2 population 
resulting from a cross between the Ditylenchus africanus resistant PC254K1 and 
susceptible Sellie. Markers are indicated on the right of each bar and the 
distances between each marker are indicated in centi Morgan (cM) on the left of 
each bar. 
105 
Eight markers mapped to LG1, six to LG2, seven to LG3, two to LG4, three to LG5, 
six to LG6 and eight to LG7. Forty-four markers were linked with the resistance trait. 
LG1 covered a relative genetic distance of 223.7 cM, LG2 a distance of 80.6 cM, LG3 
a distance of 140.2 cM, LG4 a distance of 15.5 cM, LG5 a distance of 64.8 cM, LG6 
a distance of 206.4 cM and LG7 a distance of 197.3 cM. A total relative genetic 
distance of 928.5 cM was mapped on the seven linkage groups. 
Results from the linear regression analysis on the DNA markers are shown in Table 
6.4. Association between the marker and D. africanus resistance trait was considered 
significant if the probability was P < 0.05. Although TCTCGT102 (linkage group (LG) 
1), TCTGAG490 (LG3), TCTGAG414 (LG4) and TCAGAA63 (LG6) showed a 
significant association with the resistance trait none of these markers showed to be 
closely linked to the trait since R2 values were low. 
106 
Table 6.4. Linear regression analysis of DNA markers on the seven linkage groups (LG) on 
the AFLP-based genetic linkage map of groundnut. Loci were ordered using 
MAPMAKER.EXP and posistions for putative QTLs for Ditylenchus africanus 
resistance were determined using MAPMAKER.QTL. 
Marker LG D. africanus /15 g pod P-value R2 
TCTCGT102 0.037 4.9 
TCTCGT94 0.112 2.9 
TCAGAA103 0.193 2.0 
TTACT101 0.221 1.6 
TAAGGC114 0.423 0.8 
TCAGAA89 0.475 0.6 
TCAGAA83 0.624 0.2 
TCAGAA86 0.789 0.1 
TAAGGC79 2 0.402 0.8 
TAAGGC69 2 0.455 0.7 
TCTCTA154 2 0.496 0.6 
TAAGGC66 2 0.528 0.5 
TAAGGC115 2 0.778 0.1 
TCTGAG490 3 0.034 7.0 
TCTGAG354 3 0.074 5.0 
TCTGAG203 3 0.111 4.0 
TCTGAG206 3 0.460 0.9 
TCAACT235 3 0.492 0.7 
TCTGAG197 3 0.601 0.4 
TCTGAG218 3 0.700 0.2 
TCTGAG414 4 0.000 29.1 
TCAACT442 4 0.213 2.4 
TCAGAA232 5 0.147 2.4 
TCAGAA190 5 0.944 0.0 
TCAGAA355 5 0.953 0.0 
TCAGAA63 6 0.001 11.5 
TCTCTAT119 6 0.189 2.0 
TCTCTAT137 6 0.325 1.0 
TCTCTAT103 6 0.396 0.7 
TCTCTAT136 6 0.424 0.6 
TCTCGT150 6 0.468 0.6 
TCTCCG138 7 0.153 2.1 
TCTCCG198 7 0.310 1.1 
TCTCTA163 7 0.532 0.5 
TGTCCG84 7 0.519 0.4 
TGTCCG88 7 0.511 0.4 
TGTCCG141 7 0.531 0.4 
TGTCCG68 7 0.601 0.3 
TGTCCG98 7 0.606 0.3 
107 
6.4.2.3 QTL analysis 
A QTL was detected on LG 4 between markers TCAACT442 and TCTGAG414 (Fig. 
6.8). The log likelihood for this QTL was 6.97 and the variance for the D. africanus 
numbers explained was 58.2 % (data not shown). Two more putative QTLs were 
detected on LG 6, spanning the region between markers TCTCAT103 and 
TCTCTA119 (Fig. 6.8). The log likelihood of the one QTL between markers 
TCTCAT103 and TCAGAA63 on LG 6 were 13.03 and the percentage variance 
explained was 56.1 % (data not shown). The second QTL between markers 
TCAGAA63 and TCTCTA119 on LG 6 had a log likelihood of 12.12 and the variance 
explained for this QTL is 56.1 % (data not shown). 
LG4 LG6 
JO 15 
0.0 . TCAACT442 
. TCTGAG414 i 
D. africanus1] 
25.5 
72.8 
118.2 
148.7 
206.4 
TCTCAT136 
TCTCAT137 
- TCTCAT103 
TCTCGT150 
- TCAGAA63 
TCTCTA119 
Figure 6.8. QTL-likelihood plot of markers for Ditylenchus africanus resistance on linkage 
groups (LG) 4 & 6. Markers are indicated to the right of each LG bar and the 
distances between each marker in centi Morgan (cM) to the left of the bar. 
Likelihood plots generated by MAPMAKER.QTL are indicated in blocks. The 
putative QTLs (D. africanus 1 & 2) are indicated by solid black bars. 
6.5 DISCUSSION 
Resistance or susceptibility of the parents and F2 of the reciprocal Sellie and 
PC254K1 crosses was based on Pf of the nematodes in the pods at harvesting. This 
is where the largest proportion of a D. africanus population occurs at harvesting 
108 
(Basson ef a/., 1990) and resistance can be determined with confidence only at pod 
maturity (DeWaele ef a/., 1990). 
Although the Chi2 segregation used in the linkage analysis were in the 3:1 pattern, 
the normal distribution of the F2 plants in terms of nematode numbers on pods 
indicates that this trait was quantitatively (polygenic) inherited (Roberts, 2002) and 
does not comply with Mendelian laws for heredity (Strickberger, 1976; Klug & 
Cummings, 1994). The distribution of F2 plants in terms of Pf in their pods also 
indicates that the resistance trait was polygenic (Roberts, 2002) because this trait 
demonstrated considerable variation and could not easily be categorised into 
resistant or susceptible classes (Klug & Cummings, 1994). 
Although more polymorphisms were detected in groundnut during this study than 
those by Herselman ef a/. (2004) AFLP combined with bulk segregant analysis led to 
few informative fragments, with none that were closely linked with the D. africanus 
resistance trait. This is due to a lack of polymorphism for the markers between Sellie 
and PC254K1 due to the inherently narrow genetic base of this crop (Grieshammer & 
Wynne, 1990; Halward ef a/., 1991; He & Prakash, 1997; Hopkins ef a/., 1999; 
Subramanian ef a/., 2000; Herselman, 2003). The lack of polymorphism for the 
markers between Sellie and PC254K1 complicated identification of the location of 
QTL's associated with the resistance trait during this study. Three QTL's have, 
nevertheless, been identified during this study on two separate linkage groups, which 
suggests strongly that, however strong, the resistance trait in PC254K1 is polygenic. 
Introgression of the same level of resistance governed by polygenic genes into new 
cultivars may be difficult without the use of breeder-friendly markers, though. The 
position and proportion of linkage of the QTL marker, therefore, need to be validated 
by testing the reaction on an independent mapping groundnut population using CG7 
(Chapter 3) as one of the parents. The AFLP markers can then either be used 
directly in a breeding programme or it can be converted to SCAR's (Sequence-
Characterised Amplified Region). 
109 
CHAPTER 7 
CONCLUSIONS 
To achieve the goals set for this study a multi-disciplinary approach was taken. The 
combination of nematology, plant anatomy, plant breeding and molecular biology 
enabled us to provide a meaningful contribution to the local groundnut industry. On 
completion of this study the following objectives were reached: 
i) The breeding lines PC254K1 and CG7 were identified as resistant to 
Ditylenchus africanus. The resistance expressed by these two genotypes 
is sustainable under field conditions. 
ii) The resistance expressed by PC254K1 is effective even at high 
population densities. This genotype consistently produced yields with low 
UBS % at all Pi or Pf levels tested during this study, which supports the 
nematode data. 
iii) The resistance expressed by PC254K1 is not transferred to leaf callus 
tissue of this genotype, confirming there is no short-cut for screening for 
resistance to D. africanus. 
iv) The reproduction and damage potential of D. africanus populations from 
different localities in the groundnut-production areas of South Africa is 
similar. PC254K1 expressed resistance against all the D. africanus 
populations. This genotype sustained low nematode numbers and 
consistently produced yields with low UBS %. 
v) The mechanism of resistance expressed by PC254K1 seems to be the 
inhibition of proper development and reproduction of this nematode. 
vi) Origin of the resistance trait in PC254K1 seems to be polygenic. 
vii) Eighty-four markers were mapped but none were closely linked to the 
resistance trait in PC254K1. More success may be achieved in future 
studies using the genotype CG7 from which PC254K1 was developed. 
This study was pioneering because no worthwhile resistant groundnut sources were 
identified since the discovery of D. africanus during 1987. PC254K1 and CG7 were 
the first groundnut genotypes identified with sustainable resistance to D. africanus 
under microplot as well as field conditions. In the presence of this nematode these 
two genotypes consistently maintained low nematode numbers and produced yields 
with low UBS %. PC254K1 furthermore sustained resistance even under high 
110 
nematode population pressure. The strong characteristic of PC254K1 in terms of D. 
africanus population suppression is of particular significance because the Pi of D. 
africanus in a field is irrelevant when current cultivars are grown since this nematode 
is able to build up to damaging levels during a single growing season even from a 
relatively small Pi. PC254K1 could, therefore, be used a major source of resistance 
to D. africanus in a groundnut-breeding programme for developing resistant cultivars. 
Although PC287K5 also maintained low nematode numbers in some trials, its level of 
resistance does not seem to be as strong as that of PC254K1 or CG7 and seems not 
to be as sustainable under field conditions. However, this line could still play an 
important role in the groundnut industry particularly under conditions of lower D. 
africanus infestations. Previous studies have shown that even cultivars such as 
Kwarts that show tolerance to this nematode could play a significant role under 
certain conditions. 
Development of a D. a/r/canus-resistant cultivar will benefit to the groundnut industry 
because input costs could be reduced, which in turn would increase a producer's 
profit margin especially under dry-land conditions. 
The ultimate challenge for any breeding-programme is to release new cultivars 
through the shortest possible route. One method to theoretically reduce the period of 
evaluations of groundnut genotypes against D. africanus is the use of callus tissue 
cultured from the leaves of genotypes to be screened. Unfortunately it was confirmed 
that the resistance expressed by PC254K1 in microplot and field trials was not 
transferred to callus tissue. The exercise did, however, confirm the suitability of 
groundnut leave-callus tissue cultures for propagation of D. africanus. 
The reproduction and damage potential of D. africanus populations from different 
geographically-isolated localities in the groundnut-production areas of South Africa 
was similar. Resistance of PC254K1 to all of the populations tested was confirmed. 
These results indicate that the resistant trait of a D. africanus cultivar developed from 
PC254K1 should be sustainable over the whole groundnut production area of South 
Africa. 
The absence of D. africanus from pod tissue of PC254K1 confirmed the genotype's 
resistance and suggested that the mechanism of resistance may be the inhibition of 
proper development of this nematode. The few D. africanus individuals observed 
near vascular bundles were not able to affect dissolving of the cell walls, which is 
111 
necessary for proper development and reproduction of the nematode. D. africanus is, 
therefore, not able to develop and build up to damaging population levels on 
PC254K1. The presence of low nematode numbers also means, however, that 
PC254K1 is not immune to this nematode. When PC254K1 would be rotated with 
susceptible genotypes these low nematode population levels may reach damaging 
proportions on the susceptible genotype over time. Additional control measures 
would still need to be applied. 
Information on the mechanism of resistance is important to groundnut breeders. 
Combining the resistance in PC254K1 with that of a resistant genotype from a 
different source or with other desirable characteristics could produce a cultivar of 
superior preference in the groundnut industry. 
The resistance trait in PC254K1 is seemingly governed by a number of genes. On 
the one hand this implies that resistance of PC254K1 will be more durable under 
constant selection pressure of D. africanus populations. However, although markers 
associated with the resistance trait were mapped, they were not closely linked to the 
resistance trait because of the lack of polymorphism between the parents. Three 
putative QTL's were identified but markers associated with the resistance trait need 
to be refined and developed into breeder-friendly markers that could be used in 
marker-assisted selection. These results cast doubt on the inheritance of this 
resistance, at least at the level expressed throughout the study. 
CG7 is one of the parents of PC254K1 and was tested only in one part of the study 
but there are strong indications that this genotype may have superior levels of 
resistance to D. africanus that may even surpass that of PC254K1. Identification of 
markers closely associated with the resistance trait might, therefore, be more 
successful using CG7 in stead of PC254K1. The quest for useful markers should be 
continued not only to expedite introgression of this valuable source of D. africanus 
resistance but also to ascertain that the level of resistance is sustained in commercial 
cultivars resulting from a resistance-breeding programme. 
112 
CHAPTER 8 
BIBLIOGRAPHY 
ABAWI, G.S. & MAI, W.F. 1980. Effects of initial population densities of Heterodera 
shachtii on yield of cabbage and table beets in New York State. Phytopathology 
70:481-485. 
ABDEL-MOMEN, S.M., SIMPSON, C.E. & STARR, J.L 1998. Resistance of 
interspecific Arachis breeding lines to Meloidogyne javanica and an undescribed 
Meloidogyne species. Journal of Nematology 30:341-346. 
AGRIOS, G.N. 2005. Plant Pathology. V edition, Elsevier Academic Press, California, 
USA. pp 922. 
AGUDELO, P., ROBBINS, R.T., STEWARD, J. McD. & SZALANSKI, A.L. 2005. 
Intraspecific variability of Rotylenchulus reniformis from cotton-growing regions in the 
United States. Journal of Nematology 37:105-114. 
ALTIERI, M.A. 1984. Pest management technologies for peasants: a farming system 
approach. Crop Protection 3:87-94. 
ANON. 1997. Agricultural Product Standards Act No. 119 of 1990 of the Republic of 
South Africa. Government Gazette 18056. Cape Town, South Africa, Juta & Co. pp. 
27. 
BARKER, K.R. 1991. Rotation and cropping systems of nematode control: the North 
Carolina experience-introduction. Journal of Nematology 23:342-343. 
BARKER, K.R., HUSSEY, R.S., KRUSBERG, L.R., BIRD, G.W., DUNN, R.A., 
FERRIS, H., FERRIS, V.R., FRECKMAN, D.W., GABRIEL RIDDLE, D.L, 
ROBERTS, P.A. & SCHMITT, D.P. 1994. Plant and soil nematodes: societal impact 
and focus for the future. Journal of Nematology 26:127-137. 
BARKER, K.R. & NOE, J.P. 1987. Establishing and using threshold population levels. 
jn: Veetch, J.A. & Dickson, D.W. (eds) Vistas on Nematology. Hyattsville, MD, USA, 
Society of Nematologists Inc. pp. 75-81. 
113 
BARKER, K.R. & NUSBAUM, C.J. 1971. Diagnostic and advisory programs, in: 
Zuckerman, B.M., Mai, W.F. and Rhode, R.A. (eds) Plant Parasitic Nematodes, Vol. 
1. Academic Press, New York, USA, pp. 281-301. 
BARKER, K.R. & OLTHOF, T.H.A. 1976. Relationships between nematode 
population densities and crop response. Annual Review of Phytopathology 14:327-
353. 
BASSON, S., DE WAELE, D. & MEYER, A.J. 1990. An evaluation of crop plants as 
hosts for Ditylenchus. destructor isolated from peanut. Nematropica 20:23-29. 
BASSON, S., DE WAELE, D. & MEYER, A.J. 1991. Population dynamics of D. 
destructor on peanut. Journal of Nematology 23:485-490. 
BASSON, S., DE WAELE, D. & MEYER, A.J. 1992. Effect of host plant age on 
population development and pathogenicity of Ditylenchus destructor on peanut. 
Journal of Nematology 24:310-314. 
BASSON, S., DE WAELE, D. & MEYER, A.J. 1993. Survival of Ditylenchus 
destructor in soil, hulls and seeds of groundnuts. Fundamental and Applied 
Nematology 16:79-85. 
BLOK, V.C., EHWAETI, M., FARGETTE, M., KUMAR, A., PHILLIPS, M.S., 
ROBERTSON, W.M. & TRUDGILL, D.L. 1997. Evolution of resistance and virulence 
in relation to the management of nematodes with different biology, origins and 
reproductive strategies. Nematologica 43:1-13 
BOERMA, H.R. & HUSSEY, T.S. 1992. Breeding plants for resistance to Nematodes. 
Journal of Nematology 24:242-252. 
BOLTON, C, DE WAELE, D. & BASSON, S. 1990. Comparison of two methods for 
extracting Ditylenchus destructor from hulls and seeds of groundnut. Revue de 
Nematologie 13:233-235. 
BOS, W.S. 1977. Aphelenchoides arachidis n. sp. (Nematoda: Aphelenchoidea), an 
endoparasite of the testa of groundnuts in Nigeria. Zeitschrift fur Pflanzenkrankheiten 
und Pflanzenschutz 84:95-99. 
114 
BOS, L. & PARLEVLIET, J.E. 1995. Concepts and terminology on plant/pest 
relationships: Toward consensus in Plant Pathology and Crop Protection. Annual 
Review of Phytopathology 33:69-102. 
BRIDGE, J., BOS, W.S., PAGE, L.J. & McDONALD, D. 1977. The biology and 
possible importance of Aphelenchoides arachidis, a seed-borne endoparasitic 
nematode of groundnuts from northern Nigeria. Nematologica 23:253-259. 
BRIDGE, J. & HUNT, D.J. 1985. Aphelenchoides arachidis. C.I.H. Descriptions of 
Plant-parasitic Nematodes, set 8, no. 116. London: Commonwealth Agricultural 
Bureau. 
BROWN, R.H. 1987. Control strategies in low value crops. In: Brown, R.H. & Kerry, 
B.R. (eds) Principles and practice of nematode control in crops. Sydney: Academic 
press. 
BUROW, M.D., SIMPSON, C.E., PATERSON, A.H. & STARR, J.L. 1996. 
Identification of peanut (Arachis hypogaea L.) RAPD markers diagnostic of root-knot 
nematode (Meloidogyne arenaria (Neal) Chitwood) resistance. Molecular Breeding 
2:369-379. 
CARLEY, D.H. & FLETCHER, S.M. 1995. An overview of world peanut markets. In: 
Pattee, H.E. & Stalker, H.T. (eds) Advances in Peanut Science. American Peanut 
Research and Education Society, Inc., Stillwater, USA. pp. 554-557. 
CASTILLO, M.B., MORRISON, L.S., RUSSELL, C.C & BANKS, D.J. 1973. 
Resistance to Meloidogyne hapla in peanut. Journal of Nematology 5:281-285. 
CASWELL, E.P., NELSEN, C.E. & THOMASON, I.J. 1985. A microplot method for 
recovery of entire plant root systems and their associated endoparasitic and semi-
endoparasitic nematodes. Revue de Nematologie 8:85-92. 
CHURCH, G.T, SIMPSON, C.E., BUROW, M.D. & STARR, J.L. 2001. Identification 
of RFLP loci flanking a nematode resistance locus in peanut. Proceedings of the 
American Phytopathology Society, Salt Lake City, UT. Phytopathology 91:S17'. 
115 
CHURCH, G.T., STARR, J.L. & SIMPSON, C.E. 2005. A recessive gene for 
resistance to Meloidogyne arenaria in interspecific Arachis spp. hybrids. Journal of 
Nematology 37:178-184. 
CILLIERS, A.J., SWANEVELDER, C.J., JOUBERT, H.L.N. & BARKEL, R.G. 2001. 
Catalogue of groundnut germplasm available at the ARC-Grain Crops Institute 
Potchefstroom. Fourth revised edition, pp 108. 
COLLARD, B.C.Y., JAHUFER, M.Z.Z., BROUWER, J.B. & PANG, E.C.K. 2005. An 
introduction to markers, quantitative trait loci (QTL) mapping and marker-assisted 
selection for crop improvement: The basic concepts. Euphytica 142:169-196. 
COOK, R. & EVANS, K. 1987. Resistance and tolerance. In: Brown, R.H. & Kerry, 
B.R. (eds) Principles and Practice of Nematode Control in Crops. Academic Press, 
New York. pp. 179-231. 
COOK, R. & STARR, J.L. 2006. Resistant cultivars. Inj. Perry, R.N. & Moens, M. 
(eds) Plant nemtology. CABI Publishing, pp. 392-431. 
DE WAELE, E., DE WAELE, D. & WILKEN, R. 1987. Effect of root mass on the 
efficacy of methods for extracting root-lesion nematodes from maize roots. 
Phytophylactica 19:473-474. 
DE WAELE, D., JONES, B.L., BOLTON, C. & VAN DEN BERG, E. 1989. Ditylenchus 
destructor in hulls and seeds of peanut. Journal of Nematology 21:10-15. 
DE WAELE, D., JORDAAN, E.M. & BASSON, S. 1990. Host status of seven weed 
species and their effects on Ditylenchus destructor infestation of peanut. Journal of 
Nematology 22:292-293. 
DE WAELE, D. & SWANEVELDER, C.J. 2001. Groundnut. ]n: Raemaekers, R.H. 
(ed) Crop Production in Tropical Africa, Directorate General for International Co 
operation, Brussels, Belgium, pp. 747-763. 
DE WAELE, D., VENTER, C. & MC DONALD, A.H. 1997. The peanut pod nematode 
Ditylenchus africanus. Nematology Circular No. 218. March/April, pp 6. 
116 
DE WAELE, D. & WILKEN, R. 1990. Effect of temperature on the in vitro 
reproduction of Ditylenchus destructor isolated from groundnut. Revue de 
Nematoiogie 13:171-174. 
DE WAELE, D., WILKEN, R. & LINDEQUE, J. 1991. Response of potato cultivars to 
Ditylenchus destructor isolated from groundnut. Revue Nematology 14:123-126. 
DICKSON, D.W. & DE WAELE, D. 2005. Nematode Parasites of Peanut. In: Luc, M., 
Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical 
agriculture. Second edition. CABI Publishing, pp. 393-436. 
EISENBACK, J.D. 1985. Interactions among concomitant populations of nematodes. 
In: Sasser, J.N. & Carter, W. W. (eds) An advanced treatise on Meloidogyne vol. 1. 
Biology and control. Raleigh: North Carolina State University Graphics, pp. 193-213. 
EVANS, K. & HAYDOCK, P.J.J. 1990. A review of tolerance by potato plants of cyst-
nematode attack, with consideration of what factors may confer tolerance and 
methods of assaying and improving it in crops. Annals of Applied Biology 117:703-
740. 
EVANS, K., TRUDGILL, D.L. & WEBSTER, J.M. 1993. Plant parasitic nematodes in 
temperate agriculture. CAB International, Wallingford, UK. 648 pp. 
FASSULIOTIS, G. 1979. Plant breeding for root-knot nematode resistance. ]rr 
Lambert, F. & Taylor, C.E. (eds) Root-knot Nematodes (Meloidogyne species) 
Systematics, Biology and Control. Academic Press, New York. pp. 425-453. 
FERRIS, H. 1978. Nematode economic thresholds: derivation, requirements, and 
theoretical considerations. Journal of Nematology 10:341-349. 
FERRIS, H. 1985. Density-dependent nematode seasonal multiplication rates and 
overwinter survivorship: a critical point model. Journal of Nematology 17:93-100. 
FOURIE, H. 2005. In vivo evaluation of resistance to Meloidogyne incognita race 2 
(Nematoda: Tylenchida) and identification of genetic markers for this trait in soybean 
(Glycine max), Ph.D Thesis, Katholieke Universiteit Leuven, Belgium. 
117 
FOURIE, H., Mc DONALD, A.H & LOOTS, G.C. 1999. Host suitability of South-
African commercial soybean cultivars to two root-knot nematode species. African 
Plant Protection 5:119-124. 
FOURIE, H., MIEMIE, C.M.S., Mc DONALD, A.H. & DE WAELE, D. 2008. 
Idenrification and validation of genetic markers associated with Meloidogyne 
incognita race 2 resistance in soybean, Glycine max (L.) Merr. Nematology 10:651-
661. 
FUKUDA, H. 1997. Tracheary element differentiation. The Plant Cell 9:1147-1156. 
GREGORY, W.C., GREGORY, M.P., KRAPOVICKAS, A., SMITH, B.W. & 
YARBROUGH, J.A. 1973. jn: Peanuts: Culture and uses. American Peanut Research 
Education Association. Stillwater, United Kingdom. 
GRIESHAMMER, U. & WYNNE, J.C. 1990. Isozyme variability in mature seeds of 
U.S. peanut cultivars and collections. Peanut Science 18:72-75. 
HALWARD, T.M., STALKER, H.T., LARUE, E.A. & KOCHERT, G. 1991. Genetic 
variation detectable with molecular markers among unadapted germplasm resources 
of cultivated peanut and related wild species. Genome 34:1061-1066. 
HAMMONS, R.O. 1982. Origin and early history of the peanut. In: Pattee, H.E & 
Young, C.T. (eds) Peanut science and technology. American Peanut Research and 
Education Society, Inc., Yoakum, Texas, USA. pp. 1-20. 
HAYDOCK, P.J., WOODS, S.R., GROVE, I.G. & HARE, M.C. 2006. Chemical control 
of nematodes. In; Perry, R.N. & Moens, M. (eds) Plant nemtology. CABI Publishing. 
pp. 392-431. 
HE, G. & PRAKASH, C.S. 1997. Identification of polymorphic DNA markers in 
cultivated peanut (Arachis hypogaea L.) Euphytica 97:143-149. 
HEAGLE, A.S. 1997. Abiotic diseases, in; Kokalis-Burelle, N., Porter, D.M., 
Rodriguez-Kabana, R., Smith, D.H. & Subrahmanyam, P. (eds) Compedium of 
peanut diseases. 2nd edition. St Paul MN, USA, APS Press, pp. 63-64. 
118 
HERSELMAN, L. 2003. Genetic variation among Southern African cultivated peanut 
(Arachis hypogaea L.) genotypes as revealed by AFLP analysis. Euphytica 133:319-
327. 
HERSELMAN, L, THWAITES, R., KIMMINS, F.M., COURTOIS, B., VAN DER 
MERWE, P.J.A. & SEAL, S.E. 2004. Identification and mapping of AFLP markers 
linked to peanut (Arachis hypogaea L.) resistance to the aphid vector of groundnut 
rosette disease. Theoretical and Applied Genetics 109:1426-1433. 
HINDS, M.J., SINGH, B. & ANDERSON, J.C. 1992. Determination of pod and crop 
maturity for peanuts using percent pod-fill. Canadian Journal of Plant Science 
72:1057-1065. 
HOOPER, D.J. & SOUTHEY, J.F. 1982. Ditylenchus, Anguina and other related 
genera, in: Southey, J.F. (ed) Plant nematology. Ministry of Agriculture, Fisheries and 
Food. London: Her Majesty's Stationary Office. 
HOPKINS, M.S., CASA, A.M., WANG, T., MITCHELL, S.E., DEAN, R.E., KOCHERT, 
G.D. & KRESOVICH, S. 1999. Discovery and characterization of polymorphic simple 
sequence repeats (SSRs) in peanut. Crop Science 39:1243-1247. 
HOUGH, A. & THOMASON, I.J. 1975. Effects of aldicarb on the behaviour of 
Heterodera schachtii and Meloidogyne javanica. Journal of Nematology 7:221-229. 
HUSSEY, R.S. & JANSSEN, G.J.W. 2002. Root-knot nematodes: Meloidogyne 
species. In; Starr, J.L., Cook, R. & Bridge, J. (eds) Plant resistance to parasitic 
nematodes. Egham, UK: CABI Publishing. 
ISLEIB, T.G., WYNNE, J.C. & NIGAM, S.N. 1994. Groundnut breeding, ini Smartt, J. 
(ed) The groundnut crop. Chapman & Hall, Bury St Edmunds, UK. pp. 552-623. 
JENKINS, W.R. 1964. A rapid centrifugal-flotation technique for separating 
nematodes from soil. Plant Disease Reporter 48:692. 
JOHNSON, J.T., RICH, J.R. & BOATRIGHT, A.W. 1981. A technique for establishing 
microplots in the field. Journal of Nematology 13:233-235. 
119 
JONES, F.G.W. 1985. Modelling multigenic resistance to potato cyst nematodes. 
OEPP/EPPO Bulletin 15:155-166. 
JONES, B.L. & DE WAELE, D. 1988. First report of Ditylenchus destructor in pods 
and seeds of peanut. Plant Disease 72:453. 
JONES, B.L. & DE WAELE, D. 1990. Histopathology of Ditylencus destructor on 
peanut. Journal of Nematology 22:268-272. 
KAHN, M.R. Plant nematodes. Methodology, Morphology, Systematics, Biology and 
Ecology. 
KASHA, K.J. 2008. Biotechnology and world food supply. Genome 42:642-645. 
Science Publishers, Enfield, MH, USA. pp360. 
KIM, D.G. & FERRIS, H. 2002. Relationship between crop losses and initial 
population densities of Meloidogyne arenaria in winter-grown oriental melon in Korea. 
Journal of Nematology 34:43-49. 
KLUG, W.S. & CUMMINGS, M.R. 1994. Modification of Mendelian Ratios. Jn: Starr, 
J.L., Cook, R. & Bridge, J. Concepts of genetics. Macmillan Publishing Company, 
New Jersey, USA. 
KNAUFT, D.A., GORBET, D.W. & NORDEN, A.J. 1988. Yield and market quality of 
seven genotypes grown without leaf spot control. Peanut Science 15:9-13. 
KNAUFT, D.A. & WYNNE, J.C. 1995. Peanut breeding and genetics. Advances in 
Agronomy 35:393-445. 
KOEMNING, S.R. 2000. Density-dependent yield of Heterodera g/yc/nes-resistant 
and -susceptible cultivars. Journal of Nematology 32:502-507'. 
KOKALIS-BURELLE, N., PORTER, D.M., RODRIGUEZ-KABANA, R., SMITH, D.H. 
& SUBRAHMANYAM, P. 1997. Compendium of peanut diseases. 2nd edition. St Paul 
MN, USA, APS Press, pp. 94. 
120 
KOTCON, J.B., LORIA, R. & WIXTED, D.J. 1987. Pratylenchus penetrans population 
dynamics on three potato cultivars. Journal of Nematology 19:361-368. 
KRAPOVICAS, A. & GREGORY, W.C. 1994. Taxonomy of the genus Arachis 
(Leguminosae). Bonplandia (Corrientes) 8:1-186. 
KRUSBERG, L.R. & BABINEAU, D.E. 1977. Application of plant tissue culture to 
plant nematology. In; Sharp, W.R., Larsen, P.O., Paddock, E.F. & Raghavan, V. 
(eds) Plant cell and tissue culture. Columbus: Ohio State University Press. 
LABUSCHAGNE, H., KOTZE, J. M. & WEHMER, F.C. 1980. Thielaviopsis basicola 
infection of groundnuts in South Africa. Phytophylactica 12:177-180. 
LANDBOUWEEKBLAD. 2004. Markneigings. 161 pp. 
LAMDBOUWEEKBLAD. 2005. Markneigings. 169 pp. 
LANDER, E.S., GREEN, P., ABRAHAMSON, J., BARLOW, A., DALY, M.J., 
LINCOLN, S.E. & NEWBURG, L. 1987. MAPMAKER: an interactive computer 
package for constructing primary genetic linkage maps of experimental and natural 
populations. Genomics 1:174-181. 
LILLIE, R.D. & FULLMER, H.M. 1976. Histopathologic technique and practical 
histochemistry. Fourth edition. Mc Graw-Hill Book Company, New York, U.S.A. 
LINCOLN, S., DALY, M.J. & LANDER, E. 1992. Constructing genetic linkage maps 
with MAPMAKER/EXP 3.0. Whiteheat Institute technical report, 3rd edition, 
Cambridge, Massachusetts, United States of America. 
LUC, M., BRIDGE, J. & SIKORA, R.A. 2005. Reflections on Nematology in 
Subtropical and Tropical Agriculture. ]rr Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant 
parasitic nematodes in subtropical and tropical agriculture. Second edition. CABI 
Publishing, pp. 1-10. 
MARIN, D.H., BARKER, K.R., KAPLAN, D.T., SUTTON, T.B. & OPPERMAN, C.H. 
1999. Aggressiveness and damage potential of Central American and Carribbean 
populations of Radopholus spp. in banana. Journal of Nematology 31:377-385. 
121 
Mc DONALD, A.H. 1998. Management of Ditylenchus africanus (Nematoda: 
Tylenchidae) on groundnut (Arachis hypogaea). Ph.D Thesis, Potchefstroom 
University for Christian Higher Education, Potchefstroom, South Africa. 
Mc DONALD, A.H., LOOTS, G.C, FOURIE, H. & DE WAELE, D. 2005. A microplot 
study on Ditylenchus africanus population densities and damage symptoms on 
groundnut in relation to commercial yields. Nematology 7:647-653. 
MCDONALD, D., REDDY, D.V.R., SHARMA, S.B., MEHAN, V.K. & 
SUBRAHMANYAM, P. 1998. Diseases of groundnut. Jn: Allen D.J. & Lenne J.M. 
(eds) The pathology of food and pasture legumes. CAB International, Oxon, UK. pp. 
63-124. 
McSORLEY, R. 1998. Population dynamics, jn: Barker, K.R., Pederson, G.A. & 
Windham, G.L. (eds) Plant-Nematode Interactions. American Society of Agronomy, 
Madison, Wisconsin, pp. 109-133. 
MEHAN, V.K.; MAYEE, CD., BRENNEMAN, T.B. & McDONALD, D. 1995. Stem and 
pod rots of groundnut. ICRISAT information bulletin 44:28. ICRISAT, India. 
MEYER, A.J. 1984. Die in vitro - aseptiese massateling van Pratylenchus zeae 
(Nematoda:Pratylenchinae). Phytophylactica 16:259-261. 
MIENIE, CM., LIEBENBERG, M.M.; PRETORIUS, Z.A. & MIKLAS, P.N. 2005. 
SCAR markers linked to the common bean rust resistance gene Ur-13. Theoretical 
and Applied Genetics 111:972-979. 
MURANT, A.F., ROBINSON, D.J. & GIBBS, M.J. 1995. Genus Umbravirus. ]n: 
Murphy, F.A., Fauquet, CM., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W., Martelli, 
G.P., Mayo M.A. & Summers M.D. (eds) Virus taxonomy - Classification and 
Nomenclature of Viruses: Sixth Report of the International Committee on the 
Taxonomy of Viruses. Springer-Verlag, Vienna, pp. 388-391. 
NAIDU, R.A., KIMMINS, F.M., DEOM, C, SUBRAHMANYAM, P., CHIYEMBEKEZA, 
A.J. & VAN DER MERWE, P.J.A. 1999. Groundnut Rosette. A virus disease affecting 
groundnut production in sub-Saharan Africa. Plant Disease 83:700-709. 
122 
NEL, A., KRAUSE, M. & KHELAWANLALL, N. 2007. A guide for the control of plant 
pests. Department of agriculture, Republic of South Africa. 40th edition, pp 275. 
NIBLACK, T., HUSSEY, R.S. & BOERMA, H.R. 1986a. Effects of environments, 
Meloidogyne incognita inoculum levels, and Glycine max cultivar on root-knot 
nematode-soybean interactions in field microplots. Journal of Nematology 18:338-
346. 
NIBLACK, T., HUSSEY, R.S. & BOERMA, H.R. 1986b. Effects of Heterodera 
glycines and Meloidogyne incognita on early growth of soybean. Journal of 
Nematology 18:444-450. 
NOE, J.P. 1992. Variability among populations of Meloidogyne arenaria. Journal of 
Nematology 24:404-414. 
NOE, J.P. 1998. Crop and nematode-management systems. \jv Barker, K.R., 
Pederson, G.A. & Windham, G.L. (eds) Plant and Nematode Interactions. American 
Society of Agronomy, Inc., Madison, Wisconsin, pp. 159-171. 
NUSBAUM, C.J. & FERRIS, H. 1973. The role of cropping systems in nematode 
population management. Annual Review of Phytopathology 11:423-440. 
O'BRIEN, T.P. & MC CULLEY, M.E. 1981. The study of plant structure: Principles 
and selected methods. Thermarcarphy Pty., Ltd., Melbourne, Australia, pp 59-62. 
OGALLO, J.L, GOODELL, P.B., ECKERT, J.W. & ROBERTS, P.A. 1999. 
Management of root-knot nematodes with resistant cotton cv. NemX. Crop Science 
39:418-421. 
ORTIZ, R. 1998. Critical role of plant biotechnology for the genetic improvement of 
food crops: Perspectives for the next millennium. Electronal Journal of Biotechnology 
(online) 1: Issue of August 15. 
PATERSON, A.H., LANDER, E.S., HEWITT, J.D., PETERSON, S., LINCOLN, S.E. & 
TANKSLEY, S.D. 1988. Resolution of quantitative traits into Mendelian factors by 
using a complete linkage map of restriction fragment length polymorphisms. Nature 
335:721-726. 
123 
PEREZ, E.E., MUELLER. J.D. & LEWIS, S.A. 2003. Seasonal population densities of 
Hoplolaimus columbus and their impact on soybean yield. Nematropica 33:197-207. 
PORTER, D.M., SMITH, D.H. & RODRIGUEZ-KABANA, R. 1982. |n: Pattee, H.E. & 
Young C.T. (eds) Peanut Science and Technology. American Peanut Research 
Education Society, Yoakum, Texas, United States of America, pp. 326-410. 
PRINSLOO, G.C. 1980. Thielaviopsis basicola associated with a pod disease of 
groundnuts in South Africa. Phytophylactica 12:177-180. 
REDDY, D.V.R. 1991. Groundnut viruses and virus diseases: Distribution, 
identification and control. Review of Plant Pathology 70:665-678. 
RIBAUT, J-M. & HOISINGTON, D. 1998. Marker-assisted selection: New tools and 
strategies. Trends in Plant Science 3:236-239. 
RIEKERT, H.F. 1995. An adapted method for extraction of root-knot nematode eggs 
from maize root samples. African Plant Protection 1:41-43. 
ROBERTS, P.A. 1990. Resistance to nematodes: Definitions, concepts and 
consequneses. Irr Starr, J.L. (ed) Methods for evaluating plant species for resistance 
to plant-parasitic nematodes. Society of Nematologists, United States of America. 
ROBERTS, P.A. 2002. Concepts and consequences of resistance, jni Starr, J.L., 
Cook, R. & Bridge, J. (eds) Plant resistance to parasitic nematodes. CABI Publishing, 
UK. pp. 23-42. 
ROBERTS, P.A. & MAY, D.M. 1986. Meloidogyne incognita resistance 
characteristics in tomato genotypes developed for processing. Journal of Nematology 
18:353-359. 
RODRIGUEZ-KABANA, R. 1992. Cropping systems for the management of 
phytonematodes. hr Gommers, F.J. & Maas, P.W.T. (eds) Nematology from 
Molecule to Ecosystem. European Society of Nematologists. pp. 219-233. 
SA see SOUTH AFRICA 
124 
SAGHAI-MAROOF, M.A., SOLIMAN, K.M., JORGENSEN, R.A. & ALLARD, R.W. 
1984. Ribosomal DNA spacer-length polymorphism in barley: Mendelian inheritance, 
chromosomal location, and population dynamics. Proceedings of the Natural 
Acadamy of Science USA 81:8014-8018. 
SAKHUJA, P.K. & SETHI, C.L. 1985. Screening of groundnut germplasm for 
resistance to rootknot nematode, Meloidogyne javanica. Indian Journal of 
Nematology 15:129-130. 
SEINHORST, J.W. 1957. Ditylenchus races, pathogenicity, and ecology. 
Proceedings S-19, Workshop Phytonematodes, Nashville, United States of America. 
1-7. 
SEINHORST, J.W. 1965. The relation between nematode density and damage to 
plants. Nematologica 11:137-144. 
SHARMA, S.B. & McDONALD, D. 1990. A world list of plant parasitic nematodes 
associated with groundnuts. International Arachis Newsletter 7:13-18. 
SIKORA, R.A., BRIDGE, J. & STARR, J.L. 2005. Management practices: an 
overview of integrated nematode management technologies. In; Luc, M., Sikora, R.A. 
& Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. 
Second edition. CABI Publishing, pp. 793-825. 
SIMMONDS, N.W. 1991. Genetics of horizontal resistance to disease of crops. 
Biological Review 66:189-419. 
SIMPSON, C.E., KAPROVICKAS, A. & VALLS, J.F.M. 2001. History of Arachis 
including evidence of A. hypogaeae L. progenitors. Peanut Science. 28:78-80. 
SIMPSON, C.E. & STARR, J.L. 2001. Registration of 'COAN' peanut. Crop Science 
41:918. 
SIMPSON, C.E., STARR, J.L., CHURCH, G.T., BUROW, M.D. & PATERSON, A.H. 
2003. Registration of 'NemaTAM' peanut. Crop Science 43:1561. 
125 
SMARTT, J. 1994. The groundnut farming systems and the rural economy - a global 
view. In: Smartt, J. (ed) The groundnut Crop: A Scientific Basis for Improvement. 
Chapman & Hall, London, United Kingdom, pp. 664-699. 
SMITH, O.D., BOSWELL, T.E. & THAMES, W.H. 1978. Lesion nematode resistance 
in peanuts. Crop Science 18:1008-1011. 
SOUTH AFRICA. Department of Agriculture and Water Supply. 1990. 
Ontwikkelingsprogram vir grondbone. Olie- en ProteTensadesentrum, 
Navorsingsinstituut vir Graangewasse, Potchefstroom. 
SREE LATHA, M., SHARMA, S.B., GOUR, T.B. & REDDY, D.D.R. 1998. 
Comparison of two populations of Meloidogyne javanica based on morphology, 
biology and pathogenicity to groundnut. Nematologia Mediterranea 26:189-194. 
STALKER, H.T. 1997. Peanut (Arachis hypogaea L.). Field Crops Research 53:205-
217. 
STALKER, H.T. & MOSS, J.P. 1987. Speciation, cytogenetics and utilization of 
Arachis species. Advances in Agronomy 41:1-40. 
STARR, J.L. 1984. Expression of resistance in peanuts, Arachis hypogaea, to 
Pratylenchus brachyurus: impact on screening for resistance. Journal of Nematology 
16:404-406. 
STARR. J.L., BRIDGE, J. & COOK, R. 2002. Resistance to Plant-parasitic 
Nematodes: History, Current Use and Future Potential. In; Starr, J.L., Cook, R. & 
Bridge, J. (eds) Plant resistance to parasitic nematodes. CABI Publishing, UK. pp. 1-
22. 
STRICKBERGER, M.W. 1979. Quantitative inheritance. In: Strickberger, M.W. 
Genetics 2nd edition. Collier Macmillan Publishers, London, pp 275-290. 
SUBRAMANIAN, V., GURTU, S., BAGESWARA RAE, R.C. & NIGAM, S.N. 2000. 
Identification of DNA polymorphism in cultivated groundnut using random amplified 
polymorphic DNA (RAPD) assay. Genome 43:656-660. 
126 
SUBRAHMANYAM, P., GHAMEKAR, A.M., NOLT, B.L., REEDY, D.V.R. & 
McDONALD, D. 1983. Resistance to groundnut diseases in wild Arachis species. 
Proceedings of an International Workshop on Cytogenetics of Arachis, 31 October-
2 November 1983, ICRISAT Centre, Patancheru, India. 
SUBRAHMANYAM, P., REDDY, D.V.R., SHARMA, S.B., MEHAN, V.K. & 
McDONALD, D. 1990. A world list of Groundnut diseases. Legumes Pathology 
Progress Report, 12. ICRISAT, Patancheru, India, pp. 14. 
SUMNER, M.E., KVIEN, C.S., SMAL, H. & CSINOS, A.S. 1988. On the calcium 
nutrition of peanut (Arachis hypogaea L.). I. Operation model. Journal of Fertilizer 
Issues 5:97-102. 
SVAMV, P.M. & NARASIMHAREDDY, S. 1977. Changes in the leakage of 
electrolytes from groundnut seeds (Arachis hypogaea) during after-ripening. Seed 
Science and Technology 5:654-648. 
SWANEVELDER, C.J. 1997. Grondbone - Altyd 'n wenner. Tegniese publikasie. 
Landbounavorsingsraad-lnstituut vir Graangewasse, Potchefstroom. pp. 72. 
SWART, A. & JONES, B.L. 1994. Interaction between Ditylenchus destructor and the 
nematophagous fungus Arthrobotrys dolioformis. hv Preceedings of the 32nd Annual 
Congress of the South African Society of Plant Pathology, 23-26 January, 1994, 
Christiana, Republic of South Africa, pp 126. 
TAMULONIS, J.P., LUZZI, B.M., HUSSEY, R.S., PAROTT, W.A. & BOERMA. H.R. 
1997. DNA marker analysis of loci conferring resistance to peanut root-knot 
nematode in soybean. Theoretical and Applied Genetics 95:664-670. 
TAYLOR, S.P., HOLLAWAY, G.J. & HUNT, C.H. 2000. Effect of field crops on 
population densities of Pratylenchus neglectus and P. thornei in Southeastern 
Australia; Part 1: P. neglectus. Journal of Nematology 32:591-599. 
THIES, A.J. & FERY, R.L. 2002. Host plant resistance as an alternative to Methyl 
Bromide for managing Meloidogyne incognita in pepper. Journal of Nematology 
34:374-377. 
127 
TIMPER, P., HOLBROOK, C.C. & ANDERSON, W.F. 2003. Reproduction of 
Meloidogyne spp. on resistant peanut genotypes from three breeding programmes. 
Journal of Nematology 35:417-421. 
TODD, W.J. 1986. Effects of specimen preparation on the apparent ultrastucture of 
micro-organisms. In: Aldrich, H.C. and Todd, W.J. (eds) Ultrastructure techniques for 
microorganisms. Plenum Press, New York, USA. 87 pp. 
TODD, T.C., LONG, J.H. JR. & OAKLEY, T.R. 2003. Density-dependent 
multiplication and survival rates in Heterodera glycines. Journal of Nematology 
35:98-103. 
TRUDGILL, D.L. 1986. Concepts of resistance, tolerance and susceptibility in relation 
to cyst namtodes. ]n: Lamberti, F. & Taylor, C.E. (eds) Cyst nematodes. New York: 
Plenum Press, pp 179-189. 
TRUDGILL, D.L. 1991. Resistance and tolerance of plant parasitic nematodes in 
plants. Annual Review of Phytopathology 29:167-192. 
VAN ARK, H. 1981. Eenvoudige biometriese tegnieke en proefontwerpe met spesiale 
versysing na entomologiese navorsing. Wetenskaplike pamflet No. 396. Departement 
Landbou en Visserye, Republiek van Suid-Afrika. Pretoria: Staatsdrukker. 117 pp. 
VAN DER MERWE, P.J.A. & JOUBERT, H.L.N. 1992. Tolerance of groundnut 
cultivars to the potato-rot nematode (Ditylenchus destructor). Proceedings of the 
tenth South African Maize Breeding Symposium, 1992. Technical communication 
238. pp. 86-88. 
VAN DER MERWE, P.J.A., SUBRAHMANYAN, P., KIMMINS, F.M. & WILLEKENS, 
J. 2001. Mechanisms of resistance to groundnut rosette. International Arachis 
Newsletter 21:43-46. 
VAN DER PLANK, J.E. 1978. Genetic and Molecular Basis of Plant Pathogenesis. 
Springer Verlag, Berlin. 
128 
VAN DER WALT, P.C.W. & DE WAELE, D. 1989. Mass culture of the potato rot 
nematode Ditylenchus destructor on groundnut callus tissue. Phytophylactica 21:79-
80. 
VENTER, C, DE WAELE, D. & MEYER, A.J. 1991. Reproductive and damage 
potential of Ditylenchus destructor on peanut. Journal of Nematology 23:12-19. 
VENTER, C, DE WAELE, D. & MEYER, A.J. 1993. Reproductive and damage 
potential of Ditylenchus destructor on six peanut cultivars. Journal of Nematology 25: 
59-62. 
VENTER, C, DE WAELE, D. & VAN EEDEN, C.F. 1992. Plant-parasitic nematodes 
on field crops in South Africa. 4. Groundnut. Fundamental and Applied Nematology 
15: 7-14. 
VENTER, C, VAN ASWEGEN, G., MEYER, A.J. & DE WAELE, D. 1995. Histological 
studies of Ditylenchus africanus within peanut pods. Journal of Nematology 27:284-
291. 
VIAENE, N., COYNE, D.L. & KERRY, B.R. 2006. Biological and cultural 
management. \jx Perry, R.N. & Moens, M. (eds) Plant Nematology. CABI Publishing. 
pp. 392-431. 
VOS, P., HOGERS, R., BLEEKER, M., REIJANS, M., VAN DE LEE, T., HORNES, 
M., FRIJTERS, A., POT, J., PELEMAN, J., KUIPER, M. & ZABEAU, M. 1995. AFLP: 
a new tehchnique for DNA fingerprinting. Nucleic Acids Research 21:4407-4414. 
WEAVER, D.B, RODRIGUEZ-KABANA, R. & ROBERTSON, D.G. 1985. 
Performance of selected soybean cultivars in a field infested with mixtures of root-
knot, soybean cyst, and other phytonematodes. Agronomy Journal 77:249-253. 
WEBSTER, J.M & LOWE, D. 1966. The effect of the synthetic plant-growth 
substance 2.4-dichlorophenoxyacetic acid, on the host-parasite relationships of some 
plant-parasitic nematodes in monoxenic callus culture. Parasitology 56:313-322. 
129 
WENDT, K.R. 1992. Taxonomic differentiation of three species of Ditylenchus 
(Nematoda) and their races using ribosomal DNA analysis. M.Sc Thesis, Simon 
Fraser University, Burnaby, Canada, pp. 74. 
WENDT, K.R., SWART, A., VRAIN, T.C. & WEBSTER, J.M. 1995. Ditylenchus 
africanus sp. n. from South Africa; a morphological and molecular characterization. 
Fundamental and Applied Nematology 18:241 -250. 
WENDT, K.R., VRAIN, T.C. & WEBSTER, J.M. 1993. Separation of three species of 
Ditylenchus and some host races of D. dipsaci using RFLP's. Journal of Nematology 
25:555-563. 
WENDT, K.R. & WEBSTER, J.M. 1992. Variation in the ribosomal DNA of species 
and races of Ditylenchus. Journal of Nematology 24:625. 
WINDHAM, C.L. & WILLIAMS, W.P. 1987. Host suitability of commercial corn hybrids 
to Meloidogyne arenaria and Meloidogyne incognita. Journal of Nematology 19:13-
16. 
WINDHAM, G.C. & WILLIAMS, W.P. 1988. Reproduction of Meloidogyne javanica on 
corn hybrids and inbreds. Annals of Applied Nematology 2:25-28. 
WITTWER, S.H. 1981. The 20 crops that stand between man and starvation. Farm 
Chemicals 144:17-28. 
130