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DETERMINANTS OF ROAD TRAFFIC ACCIDENT OCCURRENCES IN LAGOS STATE: SOME LESSONS FOR NIGERIA ATUBI, AUGUSTUS .O. (PH.D) ASSOCIATE PROFESSOR DEPARTMENT OF GEOGRAPHY AND REGIONAL PLANNING DELTA STATE UNIVERSITY, ABRAKA Abstract Determinants of Road Traffic Accidents have become a major public health concern worldwide: However, unlike developed or high-income countries, many developing countries have made very little progress towards addressing this problem. Nonetheless, there are ways to strengthen the reduction of injured persons that will help to lower the toll from road traffic accidents. The purpose of this paper was to elucidate ways to accomplish these goals in the context of Lagos State, Nigeria. The result of the multiple regression analysis was 0.97. This implies that the proportion of variation in the dependent variable (i.e. road traffic accidents) explained by the independent variables (i.e. length of roads, presence of road safety and population) was 97%. The F-ratio of 276.41 was higher than the table value of 2.92 at 0.05 level of confidence. This shows that accidents significantly increase with increased length of roads (km), presence of road safety corps and population. Based on the findings recommendations were proffered on how to reduce the phenomenon of traffic accidents in Lagos State, Nigeria. INTRODUCTION The World Health Organisation (WHO) estimated that 1.17million deaths occur each year worldwide due to road traffic accidents. A breakdown of the figure indicates however, that about 70percent of the deaths occur in developing countries. The increased rate of fatal road traffic accident worldwide has been attributed to population explosion and increased motorization. Increased motorization may be characterised briefly as the “automotive revolution”, that is the motorizing of urban population especially in the developing countries. In Karachi, Pakistan the police in 1998, reported 544 deaths and 793 injuries due to RTA while ambulance records noted 343 deaths and 2,048 injuries. The rates for the year 1994 were 184 injuries and 11.2 deaths per 100,000 population (WHO, 1984). Statistics indicate that over 90 percent of traffic accident situations in Nigeria can be attributed to driver errors (Aworemi et al, 2009). Road accidents appear to occur regularly at some flash points such as where there are sharp bends, potholes and at bad sections of the highways. At such points over speeding drivers usually find it difficult to control their vehicles, which then result to fatal traffic accidents, especially at night (Atubi, 2009b). Cases of fatal road traffic accidents are reported almost daily on the major highways in Lagos State. Various categories of vehicular traffic are also involved in these fatal road traffic accidents in the state. Research in this area have focused on cases of road traffic accidents, collation of road traffic accident statistics and impact assessment of road safety campaign (OECD), 1994), Becker, 1996; C.B.N., 1997; Gozias et al, 1997 and Odero et al, 2003). At the local level research in this area are concentrated on the effects of land use and human factors on road traffic accidents (Onokala, 1995; Ogunjumo, 1995 and Omojola, 2004). Motor vehicle crashes are the leading cause of death in adolescents and young adults (Taket 1986; Mohan and Romer 1991; Smith and Barss, 1991; Feachem et al, 1992; Atubi and Onokala, 2009) and of the estimated 856,000 road deaths occurring annually worldwide, 74% are in developing countries (World Bank, 1990 and Atubi, 2000b). Dramatic increases in the proportion and absolute number of traffic facilities have been witnessed in a number of developing countries, while they decreased by more than 20% in industrialised nations (Ross et al, 1991). In both Nigeria (Oluwasanmi, 1993; Ezenwa, 1986; Atubi, 2009b, 2009e and 2010c) and Kenya (NRSC, 1992), for example, a fivefold increase in traffic related fatalities was observed over the last 30 years. African and Asian countries, with relatively low vehicle densities, are experiencing substantially higher fatality rates per 10,000 vehicles than the industrialised European and North American States (Jacobs and Sayer 1983, WHO, 1984).
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Traffic crashes also impact on the economy of developing countries at an estimated cost of 1-2% of a country’s GNP per annum, as a result of morbidity, mortality and property – related costs (Fouracre and Jacobs, 1976; Jacbos and Sayer, 1983; WHO, 1989; Jabaan, 1989a and 1990; Downing, 1991). Causes of motor vehicle crashes are multi-factorial and involve the interaction of a number of pre-crash factors that include people, vehicles and the road environment (Haddon, 1980; AMA, 1983; Stansfield et al, 1992; Robertson, 1992). Human error is estimated to account for between 64 and 95% of all causes of traffic crashes in developing countries (TRL, 1990; Atubi, 2009c). A high prevalence of old vehicles that often carry many more people than they are designed to carry, lack of safety belt and helmet use, poor road design and maintenance and the traffic mix on roads are other factors that contribute to the high rate of crashes in less developed countries. Thirty-eight studies described casualties by the category of road-user. Pedestrian fatalities were highest in 75% of the studies accounting for between 41 and 75%, followed by passengers (38-51%) in 62% of the studies. Drivers were third in 55% of the studies, and never ranked first in any country. Pedal and motorcyclists killed ranked first in India (Mohan and Bawa, 1985; Sidhu et al 1993) and Surinam (CAREC, 1987). Passengers ranked first amongst the non-fatal casualties reported in 14 studies. Pedestrians were second, with the exception of one hospital based study by Dessie and Larson (1991) in Addis Ababa that reported a very high proportion of pedestrians, accounting for 91% of all traffic casualties. Driver injuries were greater in Saudi Arabia (Bener and El-Sayyad, 1985; Ofosu et al, 1988), while motorcyclists were the most involved in Taiwan (Wu and Malison, 1990). A relatively high proportion of cyclist injuries in South East Asian countries, ranging from 39 to 63%, reflect the effects of traffic mix on the roads in the region. In Nigeria, road traffic accident situation over the last three decades has been particularly disturbing. In 1976, there were 53,897 road traffic accidents resulting in 7,717 deaths. Although in 1981, the magnitude reduced to 5,114 accidents, but the fatality increased to 10,236 which mean that there was an average of 96 accidents and 28 deaths for everyday of that year (Ogunsanya, 1991). The situation in subsequent years has not been any better. The number of people killed in road accidents between 1990 and 2005 rose from 28,253, and the fatality rate remains consistently high (Atubi, 2009c). International comparison indicates that the chance of a vehicle killing someone in Nigeria is 47 times higher than in Britain. The proportion of fatalities to injuries reported is also very high. For example, while Czech Republic has only one death in 175 accidents, France, one death in 175, South Africa, one death in 47 accidents, Nigeria has one death in 2.65 accidents (Atubi, 2010c). Atubi (2010c) examined the variation patterns of RTA in Lagos state using data for 32 years (1970-2001) and observed the number and type of vehicles involved in road traffic accidents. The study revealed that private cars, buses and taxis were more prone to accidents in Lagos State. The 16 Harmonies as for the selected Local Government Areas considered contribute above 90% of the total variance in the time series. This means that more than 90% of road traffic accidents in Lagos State could be attributed to recklessness on the part of drivers, ignorance of high way codes, over speeding etc. Also, the dominant cycles of road traffic accidents observed in this study area have periodicities of 32.00 and 16.00 years with the most dominant being 32 years. This means that the dominant and strongest road traffic accident pattern of Lagos State repeats itself every 32 years. Various road safety strategies and counter measures have been used at different stages of network development. This method of seeking to prevent road accident mainly involves conscious planning, design and operations of roads. One of the most important factors in this method is the systematic identification and treatment of hazardous locations. The main objective of the study is to develop a model necessary to identify these hazardous locations on roads commonly termed as black spots. In general, the various factors that cause accidents can be broadly categorised into road related, vehicle related and driver related. THE STUDY AREA Lagos State is a suitable case study because it hosts metropolitan Lagos, Nigeria’s major traffic centre, fastest growing city, and most heavily motorized urban area in the country. Consequently, the state has one of the highest accident and casualty rates in the country (Table 1: Federal Republic of Nigeria, 1997, p. 6). Moreover, the traffic situation in Lagos State is bad because of the absence of effective planning, vehicle-misuse, poor management, inadequate street parking, traffic congestion, delays and accidents among other contributory factors. Lagos State is situated in the South Western corner of Nigeria. This elongated state spans the Guinea Atlantic coast for over 180km, from the Republic of Benin on the west to its boundary with Ogun State in the east (figure 1), while Lagos State is the smallest in Nigeria, it has over 5 percent (i.e. 9,013,534) of the country’s estimated 140 million people (National Population Census, 2006).
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Its rate of population growth has been in excess of 9 percent per annum, or 25,000 per month or 833 per day or 34 per hours in the last decade (Lagos urban Transport Project, 2002). This population increase has been accompanied by a corresponding increase in motor vehicles and traffic accidents. However, accident rates in Lagos State are still very much on the high side compared to other states in the federation. But, fatalities and non-survival indices for the state are on the decline. This is attributable to its high level of traffic congestion (which reduces the probability of the high fatality accidents resulting from over speeding) and accessibility to good post – crash medical care in the Lagos metropolitan area.
Table 1: Spatial variations in the area, population and accident parameters among the states of Nigeria, 1970-1985
States
Area
Population (1985 estimates)
Population Density Per Sq. Km 1985
Accidents per 100,000 Population (6 yr. Ave.)
Fatal Accidents per 100,000 (6 yr. Ave.)
Fatalities per 100,000 (6 yr. Ave.)
Total Causalities per 100,000 Population 6 yr. Ave.)
(*000 sq. km)
% of Nation’s Total
Number (*000)
% of Nation’s Total
Benue-Plateau
108.4
11.02
4628.1
7.04
42.7
19.63
4.62
6.30
19.75
East-Central
22.7
2.30
8621.6
13.13
379.8
22.74
3.36
4.05
19.32
Kano
43.1
4.38
6666.0
10.13
154.7
18.50
2.81
3.61
19.61
Kwara
74.3
7.55
2777.6
4.23
37.4
32.27
5.03
6.47
34.05
Lagos
14.9
1.52
1654.0
2.52
111.1
281.39
16.00
17.30
106.82
Mid-west
39.7
4.04
2927.0
4.45
73.7
48.55
6.33
8.61
46.10
North-Central
69.0
7.10
1731.0
7.20
67.8
41.73
7.42
7.76
41.67
North-Western
313.0
31.83
9021.5
13.73
28.8
13.85
2.54
3.80
22.10
Rivers
168.4
17.12
6618.0
10.07
39.3
13.84
3.19
6.11
22.98
Western
18.2
1.85
1782.7
2.71
98.0
50.10
3.26
3.67
26.24
Nigeria
35.1
3.63
5340.2
8.13
150.0
18.00
2.11
2.60
17.16
Averages of the twelve state
75.4
7.66
10951.6
16.67
145.2
34.56
5.31
6.92
29.90
Standard deviation
983.2
100.00
65720.3
100.00
66.8
32.45
4.39
5.68
28.08
Coefficient of variation
49.60
5.17
6.41
33.81
0.98
3.61
5.75
23.81
1.43
0.70
0.90
0.70
Authors Estimates Source: The Nigerian Police.
Ikeja
Ikorodu
Ojo
Amuwo
Odofin
Epe
Ibeju/Lekki
Lagos Lagoon
Badagry
Oshodi/Isolo
Lagos Mainland
Eti-Osa
Lagos
Island
Apapa
Mushin
Alimosho
Somolu
Kosofe
Surulere
Agege
Ifako
Ijaye
OGUN STATE
REPUBLIC OF BENIN
Ajeromi
Ifelodun
G U L F O F B E N I N
OGUN STATE
Water body
L.G.A. H/Q
Int’l Boundary
State Boundary
L.G.A. Boundary
N
0
10
20
30km
Key
FIG. 1: MAP OF LAGOS STATE SHOWING THE 20 L.G.AS
SOURCE: LAGOS STATE MINISTRY OF ENVIRONMENT AND PHYSICAL PLANNING (1999)
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MATERIALS AND METHODS This study, in a broad sense, undertakes an analysis of road traffic accidents in Lagos State, Nigeria. This is an attempt to conduct a detailed investigation of accident phenomenon more than the ones that had hitherto engaged the attention of researchers. Secondary data were collected from various sources, which include the records of the Federal Road Safety Commission, the Nigerian Police, Federal Ministry of Transport, and the Federal Office of Statistics. The author, reviewed the statistics for the years 1970-2001, disaggregated by administrative councils. The records contained information on crash severity (whether fatal, serious or slight), categories of road users affected, vehicles involved, and causes of crashes (whether human error, vehicle defect, or otherwise), data and traffic environment as judged by the investigating officers. Since the police only record crashes with immediate fatalities, their data generally under estimate the true magnitude of road traffic fatalities. The data collected were analysed using both descriptive and inferential statistics. DISCUSSION OF FINDINGS In order to ascertain the factors that influence the number of reported road traffic accidents in Lagos State, a multiple regression analysis was done. The dependent variable was reported number of road traffic accidents, while the independent variables considered include length of roads (in km) in Lagos state from 1970-2001, presence of road safety corps and population (appendix 1). Besides the road safety variable which was a dummy variable, 1 for years when the road safety corps are on the road, otherwise, the rest were in numbers. Out of the three regression models tried, linear form, logarithmic form and logarithmic to the power of two forms, the linear and the logarithmic to the power of two forms proved good considering the number of significant variables, sign of the coefficients and R2 value. However, the linear form was chosen based on the Durbin-Watson statistic value of 2.118 as against that of double-log form which was 1.35. The result of the analysis shows that the R2 value was 0.967, this implies that the proportion of variation in the dependent variable (i.e. road traffic accidents) explained by the independent variables was 97%. The F-ratio of 276.41 was higher than the table value of 2.92 at 0.05 level of confidence. The choice of the three variables (i.e. length of roads, presence of road safety and population) was partly influenced by data availability. Other variables like alcohol induced accidents, over speeding and reckless driving would also have been useful but were not considered for lack of data. Table 1 shows the regression result. Table 2: Regression Results of the Factors that influence road traffic accidents in Lagos State
Independent variables
Regression coefficients
Std. Error
T-start
Remark
Length of roads (km)
10.651
3.343
3.186
S
Presence of road safety
2297.47
421.32
5.45
S
Population
4.09x10-4
0.000
4.51
S
Constant
-1099.17
767.68
-1.43
S = Significant at 0.05 level of probability From table 2, length of roads in the study area (km), presence of road safety corps and population positively and significantly influenced road traffic accidents at the 0.05 level of probability in the study area. This shows that road accidents significantly increase with increased length of roads (km), presence of road safety corps and increased population. Thus, the greater the areal extent and length of highways, the higher the number of accident occurrences in Lagos State. While it is expected that large areas are likely to have greater accident occurrences than smaller areas because to higher traffic volumes, it is, however, note worthy that the more the highways of generally high quality, the greater the number of accidents. This result confirms findings from other studies that the better the quality of the roads in Nigeria, the higher the level of road traffic accidents in the country because drivers most likely over speed on good roads, leading to more frequent and fatal accidents (Onakomaiya 1988; Filani and Gbadamosi, 2007). This finding leads to two other conclusions; first, it suggests that constructing more roads will lead to increased road traffic accidents. Since increasing the number of roads may be beneficial in other ways, especially as government increases the road network as part of increasing the nation’s infrastructure base, the benefits of such developments will have to be balanced with the expected increase in road traffic accidents. Second, the findings suggest that there were more and significant levels of road traffic accidents in the periods when road safety corps was manning the road system.
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This is because the size of Nigeria’s traffic police force is too small for its road network and driver population. Moreover, the force often stays on the major highways while most of the reported road traffic accidents occur in the densely populated and motorized municipal area. In general, the variability of road traffic accident occurrences within the study area in the 32 years period can be attributed to the variations in traffic density. This is because of improvements in the quality and density of roads has not led to appreciable changes in driving, safety provisions or stronger procedures for monitoring road users’ compliance with traffic rules and regulations across the country. Moreover, poor police practices, regional disparities in traffic police strength and inadequate facilities have limited their ability to effectively enforce traffic regulations, and by extension, their ability to be effective road safety agents. It is clear from the study that the human factor is the most potent contributor to motor vehicle accidents in Nigeria. This is not surprising because people pervade all aspects of traffic accident causation: from poor road construction, corrupt traffic law enforcement, to poor road user. The driver who disobeys road traffic regulations may be the doctor or nurse in the hospital where accident victims are treated. The law enforcement officer that has no tools to work with and display a nonchalant attitude to accident victims also contributes to their eventual death. All these make the human factor the since qua non in the development of theory and the building of models for the explanation of and solution to the problem of road accidents in Nigeria and elsewhere. A fuller understanding of the many factors which contribute to road accidents in Nigeria is needed before effective measures can be applied to alleviate the present situation. Moreover, alleviating the situation requires a coordinated approach by all stakeholders including the Police, The Federal Road Safety Commission, teacher, parents, planners, doctors and individuals members of society. IMPLICATION OF FINDINGS/RECOMMENDATIONS For Nigeria, the measures in table 3 might help to reduce the country’s high road accident rated and improve her overall road safety. Table 3: Suggested road accident prevention strategies for Nigeria
Ensuring that drivers are constantly trained to ensure that they operate vehicles safely and comply with traffic regulations at all times.
Learn from other countries with better traffic management systems e.g. the U.S. and Great Britain
Rationalise the use of traffic police check points
Ensure adequate funding for the Federal Road Safety Corps
To deal with the high accident situation in Lagos State, the authorities should: First, decongest the CBD by staggering work hours so that all workers do not leave for work at approximately the same time. This would reduce the number of workers plying the roads during peak hours and thus ease traffic congestion in the peak hours. Second, strengthen the state’s traffic patrol effort by (i) ensuring that local traffic police work closely with the Federal Road Safety Commission (FRSC), (ii) Regularly patrolling the highways and side streets and, (iii) Ensuring that local police are familiar with the temporal and spatial distribution of road traffic accidents within the their area of jurisdiction. Third, ensure that there is enough political commitment to deal with the long term issue of reducing road accidents; long term efforts to reduce the states high auto accidents rate requires (i) constant monitoring of the states roads (ii) dedicating enough resources to road safety in spite of extreme budgetary constraints, (iii) sufficient allocation of road safety research funds to identify problems in need of remedial action and to maintain up to-base data for road safety decision – making and public education and, (iv) Coordinating road safety polices at sub-regional and regional levels. Fourth, implement preventive road accident measures including proper design of road networks as well as the planning of the general public transport system to ensure that it runs in an effective and efficient manner as this would reduce the volume of the vehicles plying the roads. Fifth, consistently implement road safety measures that go beyond ad-hoc public pronouncements and traffic control activities on setting up of temporary road blocks or short-term crackdowns on commercial and private vehicles (usually after an important personality is killed in a crash). Such efforts should address broader road safety policy issues and the various underlying causes. For this to happen, it is important to enlist the necessary political good will and commitment to road traffic injury prevention.
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Sixth invest in the repair of most Lagos roads, many of which are currently in need of repair and maintenance. For this to happen the institutional machinery needed to ensure the timely construction and repair of roads should be strengthened. CONCLUSION Traffic accident injuries and fatalities are a serious problem all over the world. In Nigeria, the incidence has become a very disturbing phenomenon as the country is presently ranked among countries with the highest traffic injuries and fatalities in the world (Jacobs et al, 1986; Onakomaiya, 1991; Daramola, 2004). Moreover, the country’s road safety campaigns are inadequate and often limited to festive seasons. At other times, such campaigns are often limited to media and roadside slogans like “drive carefully”, accident kill more passengers”, “drive to stay alive”, and only living celebrates”. For these campaigns to be effective, they must be year-round and not just limited to festive seasons. REFERENCES American Medical Association (1983) Automobile-Related injuries: Components, Trends and Prevention. Journal of the American Medical Association, 249: 3216-3222. Atubi, A.O. (2009b) “Urban Transportation: An Appraisal of Features and Problems in the Nigerian Society”. International Journal of Geography and Regional Planning. Vol. 1, No. 1, Pp. 58-62. Atubi, A.O. (2009c) Modelling Road Traffic Accidents in Lagos State, South Western Nigeria. Journal of Society and State. Vol. 1, No. 1 & 2, Pp. 57-74. Atubi, A.O. (2010b) Road Transport System Management and Traffic in Lagos, South Western Nigeria. Journal of African Research Review. Vol. 4, No. 4, Pp. 459-470. Atubi, A.O. (2010c) Road Traffic Accident Variations in Lagos State, Nigeria: A Synopsis of Variance Spectra. Journal of African Research Review. Vol. 4, No. 2, Pp. 197-218. Atubi, A.O. and Onokala, P.C. (2009) Contemporary Analysis of Variability in Road Traffic Accidents in Lagos State, Nigeria. Journal of African Geographical Review. Vol. 28, Pp. 11-41. Aworemi, A. and Alawode, E.C. (2009) Roads to where? Newswatch, December 30, 17 (13) 3. Backer, W. (1996) “Impact Assessment of Road Safety Campaign” Transportation Review. 16 (4): 345-355. Bener, A. and El-Sayyad, G.M. 91985) Epidemiology of Motor Vehicle Accidents in Jeddeh. Journal of the Royal Society of Health, 105:200-201. CAREC (1987) Blood Alcohol Levels in Traffic Accident Victims, Surinam, February, 1987. CAREC Survivor Report, 13, 1-14. Central Bank of Nigeria (1997) Annual Report and Statement of Account for the Year Ended 31st December, CBN, Abuja. Daramola, A.Y. (2004) “Innovative Options for Financing Transport Infrastructure in Nigeria” In Nisereel, The Magazine of the Nigerian Institute of Social and Economic Research, Nos 4 & 5, December, Ibadan. Dessie, T. and Larson, C.P. (1991): The Occurrence and Driver Characteristics Associated with Motor Vehicle Injuries in Addis Ababa, Ethiopia. Journal of Tropical Medicine and Hygiene, 94:395-400. Downing, A.J. (1991) Driver Training of Africa: the UN-ECA Driving Manual. Proceedings of the Second African Road Safety Congress. Oslo: Institute of Transport Economics (TOI) Ezenwa, A.O. (1986) “Prevention and Control of Road Traffic in Nigeria” Journal of the Royal Society of Health. Vol. 106, No. 1, Pp. 25-26. Feachem, R.G.A,; Kjellstorm, T,; Murray, C.J.L.; Over, M. and Philips, M.A. (1992) The Health of Adults in the Developing World, London: Oxford University Press. Filani, M.O. and Gbadamosi, K.T. (2007) Spatial and Temporal Pattern of Road Traffic Accident Occurrences in Nigeria: 1970-1995. Nigerian Geographical Journal. Vol. 5, No. 1, Pp. 55-70. Fouracre, P. R. and Jacobs, G.D. (1976) Comparative Accident Costs in Developing Countries. TRRL Supplementary Report 206. Crowthorne: Transport and Road Research Laboratory. Gozias, J.C.; Matsouki, E.C. and Yannis, G.D. (1997) “An Analysis of Factors Affecting Road Safety: The Greek Experience” Journal of Transportation Geography 5 (3): 325-316. Haddon, W. (1980) Advances in the Epidemiology of Injuries as a basis for public policy. Public Health Reports. 95: pp. 411-421.
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Jacobs, G.D. and Cutting, C.A. (1986) “Further Research on Accident Rates in Developing Countries”. Accident Analysis and Prevention. Vol. 18, No. 2, pp. 119-127. Jacobs, G.D. and Sayer, I.A. (1983) Road Accidents in Developing Countries’. Transport and Road Research laboratory, TRRI Supplementary Report 897 Crowthorne. Jadaan, K.S. (1989a) Road Accidents Costs in Jordan. Journal of the Royal Society of Health, 109: 144-146. Jadaan, K.S. (1990) Traffic Accidents in Kuwait: An Economic Dimension. Accident Analysis and Prevention. 22:399-401. Mohan, D. and Bawa, P.S. (1985) An Analysis of Road Traffic Accident Fatalities in Delhi, India. Accident Analysis and Prevention, 17: 33-45. Mohan, D. and Romer, C.J. (1991) Accident Mortality and Morbidity in Developing Countries. In Manciaux M. and Romer, C.J. (ed) Accidents in Childhood and Adolescence: The Role of Research, Pp. 31-38. Geneva: World Health Organisation. National Road Safety Council of Kenya (1992) Accident Statistics, 1983-1990. Nairobi: Ministry of Public Works, Government of Kenya. Odero, W. Khayes, I.M. Meda, P. M. (2003) Road Traffic Accident in Kenya: Magnitude, Causes and Status of Intervention. Injury Control and Safety Promotion, 10, Pp. 53-61. OECD, (1994) Managing the Environment: the Role of Economic Instrument. Paris Ofosu, J.B.; Abonammoh, A.M. and Bener, A. (1988): A Study of Road Traffic Accidents in Saudi Arabia. Accident Analysis and Prevention, 20:95-111. Ogunjumo, A. (1995) En Evaluation of Nigerian Airways’ and Foreign Management Consultants’ domestic Operations: 1974-1986: The Nigerian Journal of Economic and Social Studies, Pp. 34 (1). Ogunsanya, A.A. (1991) Empirical Cases Studies of Accident and Safety Control of Mass Transit Agencies in Nigeria. In Bolade T, and Ogunsanya A. (eds) Accident Control and Safety measures in Mass Transit Operations in Nigeria. Ibadan: University Press. Pp. 87-115. Oluwasanmi, A.J. (1993) Road Accident Trends in Nigeria. Accident Analysis Prevention 25 (4) Pp. 485-487. Omojola, A.S.(2004) “Transport and Communication” in Kayode, M.O. and Usman, Y.B. (eds) Nigeria since Independence. Vol. 2, Ibadan; Heinemann. Pp. 132-152. Onakomaiya, S.O. (1988) Unsafe at any Speed: Toward Road Transportation for Survival: Inaugural Lecture, University of Ilorin, Ilorin. Onakomaiya, S.O. (1991) General Trend of Safety and Accident Records in Nigerian Transport Sector. In: Bolade T. and Ogunsanya, A. (eds) Accident Control and Safety Measures in Mass Transit Operation in Nigeria, Ibadan University Press, P. 11. Onokala, P.C. (1995) The Effect of Landuse on Road Traffic Accident in Benin-City, Nigeria. Journal of Transport Studies, Vol. 1, No. 1, Pp. 34-44. Robertson, L.S. (1992) Injury Epidemiology. Oxford University Press, New York. Ross, A. Baguley, C. Hills, V. Mchonald, M. and Silcock, d. (1991) Towards Safer Roads in Developing Countries: A guide for Planners and Engineers. Crowthorne, U.K.; Transport Research laboratories. Sidhu, D.S.; Sodi, G.S. and Banerjee, A.K. (1993) Mortality Profile in Trauma Victims. Journal of the Indian Medical Association, 1:16-18. Smith, G.S. and Barss, P. (1991) Unintentional Injuries in Developing Countries: The Epidemiology of a neglected problem. Epidemiology Review. 13, 228-266. Stansfield, S.K.; Smith, J.S. and McGreevey, W.P. (1992) Injury. In Diseases Priorities in Developing Countries. (eds) Jamison, D.T. and Mosley, W.H. World Bank, Oxford University Press, London, Pp. 609-633. Taket, A. (1986) Accident Mortality in Children, Adolescence and Young Adults. World Health Statistics Quarterly, 39:232-256. Transport and Research Laboratory (1999) Towards Safer Roads in Developing Countries, TRL, Crowthorne, U.K. World Bank (1990) “Road Safety Problem in the Third World”. The Urban Edge. Vol. 14, No. 1. World Health Organisation (1984) Road Traffic Accidents in Developing Countries, Geneva: WHO, TRS 703 World Health Organisation (1989) Analysis of Achievements of Traffic Safety in Industrialised and Developed Countries. Wu, S.I. and Malison, M.D. (1990) Motor Vehicle Injuries in Taiwan. Asia Pacific Journal of Public Health, 4:72-75.
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APPENDIX I Population, length of roads (in km), and presence of road safety corps in Lagos State from 1970-2001
Year
Population
Length of roads (km)
Road safety presence
1970
1,028,327
140
0 (No)
1971
1,085,913
142
0
1972
1,146,724
144
0
1973
1,210,940
147
0
1974
1,278,752
150
0
1975
1,350,362
152
0
1976
1,425,982
154
0
1977
1,505,836
156
0
1978
1,647,384
157
0
1979
1,802,238
157
0
1980
1,971,648
157
0
1981
2,156,982
158
0
1982
2,359,938
161
0
1983
2,581,553
164
0
1984
2,824,218
160
0
1985
3,089,694
148
0
1986
3,380,125
144
1 (yes)
1987
3,697,856
130
1
1988
4,024,554
130
1
1989
4,425,726
130
1
1990
4,842,744
130
1
1991
5,680,000
135
1
1992
6,213,920
135
1
1993
6,790,028
150
1
1994
7,097,141
194
1
1995
7,416,512
197
1
1996
7,661,256
203
1
1997
7,914,077
200
1
1998
8,175,241
250
1
1999
8,445,025
250
1
2000
8,723,709
256
1
2001
9,011,591
300
1
Source: Lagos Metropolitan Area Transport Authority (LAMATA, 2002); Master Plan Project, Urban and Regional Planning Division, Lagos State Ministry of Economic Planning and Land Matters, 2002. * There was a decline in length of roads in the country between 1985 and 1992 because these were the years of military rule in Nigeria and the various ministry regimes of that era disregarded the development of infrastructure such as roads. Moreover, flagrant abuse of power and massive corruption greatly diminished the resources available for road and other infrastructural developments.
Monday 24 September 2012
microbiological media preparation
GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.
Microbiological Media Preparation
Bacteria and fungi are grown on or in microbiological media of various types. The medium
that is used to culture the microorganism depends on the microorganism that one is trying to
isolate or identify. Different nutrients may be added to the medium, making it higher in
protein or in sugar. Various pH indicators are often added for differentiation of microbes
based on their biochemical reactions: the indicators may turn one color when slightly acidic,
another color when slightly basic. Other added ingredients may be growth factors, NaCl, and
pH buffers which keep the medium from straying too far from neutral as the microbes
metabolize.
In this exercise, you will make all-purpose media called trypticase soy broth and trypticase
soy agar. These 2 media----one a liquid and the other a solid---are the exact same formula
save for the addition of agar agar (really---agar agar), an extract from the cell walls of red
algae.
The old way to make media was by the cookbook method--- adding every ingredient bit by bit.
The only time that is done today is when making a special medium to grow a certain finicky
organism, where particular growth factors, nutrients,
vitamins, and so on, have to be added in certain amounts.
This medium is called a chemically defined medium
(synthetic). Fortunately, the most common bacteria that we
want to grow will do nicely with media that we commonly use
in lab. Some of our media is bought, but most is produced
in the prep area behind the lab. Since this type of medium
has some unknown ingredients, or sometimes unknown
quantities it is called complex media.
It is really very simple to make complex media these days:
rehydrate the powder form of the medium
stir and boil the agar medium to get the agar powder
dissolved (if making an agar medium rather than a
broth medium)
distribute the medium into tubes
autoclave to sterilize the tube media
autoclave the agar medium for plate production
and then pour into sterile petri dishes
STERILIZATION AND THE AUTOCLAVE
When microbiological media has been made, it still has to be sterilized because of microbial
contamination from air, glassware, hands, etc. Within a few hours there will be thousands
of bacteria reproducing in the media so it has to be sterilized quickly before the microbes
start using the nutrients up. The sterilization process is a 100% kill, and guarantees that the
medium will stay sterile UNLESS exposed to contaminants by less than adequate aseptic
technique to exposure to air.
Media sterilization is carried out with
the autoclave, basically a huge steam
cooker. Steam enters into a jacket surrounding
the chamber. When the pressure from the
steam is at a certain point in the jacket, a valve
allows the steam to enter the chamber. The
pressure will go up over 15 pounds per square
inch (psi): at this point the timer begins to
count down--- usually for 15 minutes, depending
on the type of media. The high pressure in a
closed container allows the temperature to go
above the highest temperature one could get
by just boiling, around 121 degrees C.
Therefore, the parameters for sterilization with
an autoclave are 121 C at >15 psi for 15
minutes. Fifteen minutes is the thermal death time for most organisms (except some really
hardy sporeformers).
The prepared media is distributed in different ways, depending on the form one is making.
Broths and agar deeps are dispensed into tubes and then sterilized. Agar slant tubes are
sterilized and then the rack is tilted to allow the agar to solidify in a slanted fashion. Agar
medium which will be poured into plates is sterilized in a flask, and then poured afterward.
Not all media or solutions can be sterilized via an autoclave. Certain high-protein solutions
such as urea, vaccines, and serum will denature in the extreme heat, and so they may have
to be filter-sterilized without heat.
OBJECTIVES:
Understand how to make media, how to sterilize it, and how to distribute it in different
formats.
Produce TSA plates, TSA slants, and TSB which will be used in subsequent lab periods.
Understand the basics of an autoclave and how it sterilizes, including parameters.
MATERIALS NEEDED: per table
1 sleeve of plastic petri plates (may need a couple more if you pour light)
1 flask melted TSA (in 50C water bath)
1 spatula 24 medium, nonsterile test tubes
2 plastic weigh boats 1 small test tube rack
1 jar nutrient trypticase soy broth powder 12 yellow caps
1 jar agar powder 12 green caps
1-1 liter Erlenmeyer flask 1 pipet pump
1 magnetic stir bar 1 pipet disposal jar
1 graduated cylinder several nonsterile glass10 ml pipets
THE PROCEDURE: (see diagram below text for the entire production)
1. Your table will pour your plates, using a liquified, sterile agar medium which has
previously been made for you (should be sitting in a water bath). The flasks of melted TSA are
in the water bath which is set to 45. This is to prevent them from solidifying, since agar
solidifies at around 42 C. Remove your flask from the water bath only when you are ready
to pour your plates.
2. Set the petri dishes out, RIGHT SIDE UP (small side down), tops covering the dishes.
When you are ready to pour your plates, obtain your flask of agar and pour the plates.
Depending on whether you pour `heavy` or pour `light', you may use anywhere between 18-
22 petri dishes. Cover the bottom of each plate approximately 2/3 of the way across, put on
the lid and gently rotate the plate to distribute the agar across the entire dish bottom. Allow
all plates to stand until they are completely solidified. Once they have solidified, place them
on the tray at the instructor's table. Agar plates are always placed UPSIDE DOWN (bottom
dish with agar on top), definitely so when you are incubating your cultures.
3. Why are agar plates incubated upside down? Two reasons:
Although there may be air contaminants in the incubator, it will be more difficult for
them to get onto plates, and then move UP onto the agar to settle since gravity is
working against this movement.
Often you will see a bit of water condensation on the petri dish cover. The water
molecules are cohesive and tend to run together. If the plate is sitting right side up,
the water droplets can fall onto the agar, creating a kind of little lake on the agar
surface, messing up the plates. Upside down plates prevents the condensation from
dropping on the agar surface.
4. Begin making the TSB (broth) by pouring
500ml of distilled water into a 1L flask. Put in the
stir bar and turn on the stir plate so that the
surface is just disturbed. Add 6.5 grams of the
TSB powder to this flask and allow it to dissolve
(will happen quickly). No heat need be applied at
this stage.
5. Once the powder is dissolved, pipet out twelve
3ml aliquots into tubes and cover each tube with a
green cap.
NOTE: Green caps are always used for TSB.
6. With the remaining solution (about 464ml) still
stirring, add 8 grams of agar powder.
7. The next step will require you to apply heat to the mixture. Before you do this,
however, you should be aware that agar has a strong tendency to boil over when it reaches
100C. Someone in your group should be watching the flask at all times once you see
steam coming off of it. At the first sign that the mix is near boiling, REMOVE it from the
hot plate (paper towels around the flask neck). DO NOT simply turn off the heat, letting
the flask sit there. The metal plate retains a significant amount of heat, and turning off the
heat will not prevent the flask from boiling over. Folded paper towels allow you to grasp
the flask neck tightly, yet not burn your hand.
8. Have you read step 7? Ok, then you can turn on the heat to setting 9 (not High). Make
sure that the magnetic bar is stirring the solution.
9. When the mixture boils and the agar dissolves, it will turn clear, deeper tan. Remove it
from the heat and pipet out twelve 6ml aliquots into medium tubes for slants (they will not be
BE slants until removed from the autoclave and tilted to the side to solidify). Pipet out four
6ml aliquots into medium tubes for agar deeps. Cover the slant tubes with yellow caps and
the deep tubes with black caps. NOTE: from this point on, yellow caps will be used for nutrient
agar slants. Also note: if the agar solidifies in the tip of the pipet, dispose of the pipet in the
pipet jar and get another one. To prevent this from happening, either pipet out all the tubes at
the same time, or leave the pipet in the flask of melted agar.
10. Place all of the tubes you have pipetted out in the plastic autoclave racks on the
instructor's table as well as the remaining of your melted agar. All agar deeps go in one rack,
broths in another rack, etc.
11. Cover the flask with the remaining agar medium (to be sterilized) with a piece of foil and
place in the metal tray on instructor’s table (for sterilization).
12. Dispose of your used pipets in the pipet holder. These glass pipets are reusable, so
don't throw them in the trash.
QUESTIONS:
1. What is a complex medium?
2. Why are pH buffers added to the growth media for microbes?
3. How can the temperature in the autoclave go above boiling temperature of 212 F?
4. Why do you have boil the agar solution BEFORE dispensing it into tubes?
5. At what temperature does agar solidify?
OLANREWAJU ISRAEL.+23470334765
Bacteria and fungi are grown on or in microbiological media of various types. The medium
that is used to culture the microorganism depends on the microorganism that one is trying to
isolate or identify. Different nutrients may be added to the medium, making it higher in
protein or in sugar. Various pH indicators are often added for differentiation of microbes
based on their biochemical reactions: the indicators may turn one color when slightly acidic,
another color when slightly basic. Other added ingredients may be growth factors, NaCl, and
pH buffers which keep the medium from straying too far from neutral as the microbes
metabolize.
In this exercise, you will make all-purpose media called trypticase soy broth and trypticase
soy agar. These 2 media----one a liquid and the other a solid---are the exact same formula
save for the addition of agar agar (really---agar agar), an extract from the cell walls of red
algae.
The old way to make media was by the cookbook method--- adding every ingredient bit by bit.
The only time that is done today is when making a special medium to grow a certain finicky
organism, where particular growth factors, nutrients,
vitamins, and so on, have to be added in certain amounts.
This medium is called a chemically defined medium
(synthetic). Fortunately, the most common bacteria that we
want to grow will do nicely with media that we commonly use
in lab. Some of our media is bought, but most is produced
in the prep area behind the lab. Since this type of medium
has some unknown ingredients, or sometimes unknown
quantities it is called complex media.
It is really very simple to make complex media these days:
rehydrate the powder form of the medium
stir and boil the agar medium to get the agar powder
dissolved (if making an agar medium rather than a
broth medium)
distribute the medium into tubes
autoclave to sterilize the tube media
autoclave the agar medium for plate production
and then pour into sterile petri dishes
STERILIZATION AND THE AUTOCLAVE
When microbiological media has been made, it still has to be sterilized because of microbial
contamination from air, glassware, hands, etc. Within a few hours there will be thousands
of bacteria reproducing in the media so it has to be sterilized quickly before the microbes
start using the nutrients up. The sterilization process is a 100% kill, and guarantees that the
medium will stay sterile UNLESS exposed to contaminants by less than adequate aseptic
technique to exposure to air.
Media sterilization is carried out with
the autoclave, basically a huge steam
cooker. Steam enters into a jacket surrounding
the chamber. When the pressure from the
steam is at a certain point in the jacket, a valve
allows the steam to enter the chamber. The
pressure will go up over 15 pounds per square
inch (psi): at this point the timer begins to
count down--- usually for 15 minutes, depending
on the type of media. The high pressure in a
closed container allows the temperature to go
above the highest temperature one could get
by just boiling, around 121 degrees C.
Therefore, the parameters for sterilization with
an autoclave are 121 C at >15 psi for 15
minutes. Fifteen minutes is the thermal death time for most organisms (except some really
hardy sporeformers).
The prepared media is distributed in different ways, depending on the form one is making.
Broths and agar deeps are dispensed into tubes and then sterilized. Agar slant tubes are
sterilized and then the rack is tilted to allow the agar to solidify in a slanted fashion. Agar
medium which will be poured into plates is sterilized in a flask, and then poured afterward.
Not all media or solutions can be sterilized via an autoclave. Certain high-protein solutions
such as urea, vaccines, and serum will denature in the extreme heat, and so they may have
to be filter-sterilized without heat.
OBJECTIVES:
Understand how to make media, how to sterilize it, and how to distribute it in different
formats.
Produce TSA plates, TSA slants, and TSB which will be used in subsequent lab periods.
Understand the basics of an autoclave and how it sterilizes, including parameters.
MATERIALS NEEDED: per table
1 sleeve of plastic petri plates (may need a couple more if you pour light)
1 flask melted TSA (in 50C water bath)
1 spatula 24 medium, nonsterile test tubes
2 plastic weigh boats 1 small test tube rack
1 jar nutrient trypticase soy broth powder 12 yellow caps
1 jar agar powder 12 green caps
1-1 liter Erlenmeyer flask 1 pipet pump
1 magnetic stir bar 1 pipet disposal jar
1 graduated cylinder several nonsterile glass10 ml pipets
THE PROCEDURE: (see diagram below text for the entire production)
1. Your table will pour your plates, using a liquified, sterile agar medium which has
previously been made for you (should be sitting in a water bath). The flasks of melted TSA are
in the water bath which is set to 45. This is to prevent them from solidifying, since agar
solidifies at around 42 C. Remove your flask from the water bath only when you are ready
to pour your plates.
2. Set the petri dishes out, RIGHT SIDE UP (small side down), tops covering the dishes.
When you are ready to pour your plates, obtain your flask of agar and pour the plates.
Depending on whether you pour `heavy` or pour `light', you may use anywhere between 18-
22 petri dishes. Cover the bottom of each plate approximately 2/3 of the way across, put on
the lid and gently rotate the plate to distribute the agar across the entire dish bottom. Allow
all plates to stand until they are completely solidified. Once they have solidified, place them
on the tray at the instructor's table. Agar plates are always placed UPSIDE DOWN (bottom
dish with agar on top), definitely so when you are incubating your cultures.
3. Why are agar plates incubated upside down? Two reasons:
Although there may be air contaminants in the incubator, it will be more difficult for
them to get onto plates, and then move UP onto the agar to settle since gravity is
working against this movement.
Often you will see a bit of water condensation on the petri dish cover. The water
molecules are cohesive and tend to run together. If the plate is sitting right side up,
the water droplets can fall onto the agar, creating a kind of little lake on the agar
surface, messing up the plates. Upside down plates prevents the condensation from
dropping on the agar surface.
4. Begin making the TSB (broth) by pouring
500ml of distilled water into a 1L flask. Put in the
stir bar and turn on the stir plate so that the
surface is just disturbed. Add 6.5 grams of the
TSB powder to this flask and allow it to dissolve
(will happen quickly). No heat need be applied at
this stage.
5. Once the powder is dissolved, pipet out twelve
3ml aliquots into tubes and cover each tube with a
green cap.
NOTE: Green caps are always used for TSB.
6. With the remaining solution (about 464ml) still
stirring, add 8 grams of agar powder.
7. The next step will require you to apply heat to the mixture. Before you do this,
however, you should be aware that agar has a strong tendency to boil over when it reaches
100C. Someone in your group should be watching the flask at all times once you see
steam coming off of it. At the first sign that the mix is near boiling, REMOVE it from the
hot plate (paper towels around the flask neck). DO NOT simply turn off the heat, letting
the flask sit there. The metal plate retains a significant amount of heat, and turning off the
heat will not prevent the flask from boiling over. Folded paper towels allow you to grasp
the flask neck tightly, yet not burn your hand.
8. Have you read step 7? Ok, then you can turn on the heat to setting 9 (not High). Make
sure that the magnetic bar is stirring the solution.
9. When the mixture boils and the agar dissolves, it will turn clear, deeper tan. Remove it
from the heat and pipet out twelve 6ml aliquots into medium tubes for slants (they will not be
BE slants until removed from the autoclave and tilted to the side to solidify). Pipet out four
6ml aliquots into medium tubes for agar deeps. Cover the slant tubes with yellow caps and
the deep tubes with black caps. NOTE: from this point on, yellow caps will be used for nutrient
agar slants. Also note: if the agar solidifies in the tip of the pipet, dispose of the pipet in the
pipet jar and get another one. To prevent this from happening, either pipet out all the tubes at
the same time, or leave the pipet in the flask of melted agar.
10. Place all of the tubes you have pipetted out in the plastic autoclave racks on the
instructor's table as well as the remaining of your melted agar. All agar deeps go in one rack,
broths in another rack, etc.
11. Cover the flask with the remaining agar medium (to be sterilized) with a piece of foil and
place in the metal tray on instructor’s table (for sterilization).
12. Dispose of your used pipets in the pipet holder. These glass pipets are reusable, so
don't throw them in the trash.
QUESTIONS:
1. What is a complex medium?
2. Why are pH buffers added to the growth media for microbes?
3. How can the temperature in the autoclave go above boiling temperature of 212 F?
4. Why do you have boil the agar solution BEFORE dispensing it into tubes?
5. At what temperature does agar solidify?
OLANREWAJU ISRAEL.+23470334765
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